A Guide to Starting and Operating a Nursery 

for Native and Traditional Plants

Tropical Nursery Manual

Kim M. Wilkinson • Thomas D. Landis • Diane L. Haase

Brian F. Daley • R. Kasten Dumroese

EDITORS

Forest 

Service

Agriculture

Handbook 732

April

2014

United States Department of Agriculture

Tropical Nursery Manual

A Guide to Starting and Operating a 

Nursery for Native and Traditional Plants

U.S. Department of Agriculture, Forest Service

Agriculture Handbook 732

April 2014

Edited by 

Kim M. Wilkinson 

Thomas D. Landis

Diane L. Haase 

Brian F. Daley

R. Kasten Dumroese
The use of trade or firm names in this publication is for 

reader information and does not imply endorsement by 

the U.S. Department of Agriculture, Forest Service of any 

product or service.

Pesticides used improperly can be injurious to humans, 

animals, and plants. Follow the directions and heed all precautions on the labels. Store pesticides in original containers under lock and key—out of the reach of children and 

animals—and away from food and feed. Apply pesticides so 

that they do not endanger humans, livestock, crops, beneficial insects, fish, and wildlife. Do not apply pesticides when 

there is danger of drift, when honey bees or other pollinating 

insects are visiting plants, or in ways that may contaminate 

water or leave illegal residues. Avoid prolonged inhalation of 

pesticide sprays or dusts; wear protective clothing and equipment if specified on the container. If your hands become 

contaminated with a pesticide, do not eat or drink until you 

have washed. In case a pesticide is swallowed or gets in the 

eyes, follow the first-aid treatment given on the label, and get 

prompt medical attention. If a pesticide is spilled on your 

skin or clothing, remove clothing immediately and wash skin 

thoroughly. Do not clean spray equipment or dump excess 

spray material near ponds, streams, or wells. Because it is 

difficult to remove all traces of herbicides from equipment, 

do not use the same equipment for insecticides or fungicides that you use for herbicides. Dispose of empty pesticide 

containers promptly. Have them buried at a sanitary landfill dump, or crush and bury them in a level, isolated place. 

NOTE: Some States have restrictions on the use of certain 

pesticides. Check your State and local regulations. Also, because registrations of pesticides are under constant review by 

the Federal Environmental Protection Agency, consult your 

county agricultural agent or State extension specialist to be 

sure the intended use is still registered.

The U.S. Department of Agriculture (USDA) prohibits 

discrimination in all its programs and activities on the 

basis of race, color, national origin, age, disability, and 

where applicable, sex, marital status, familial status, 

parental status, religion, sexual orientation, genetic information, political beliefs, reprisal, or because all or 

part of an individual’s income is derived from any public 

assistance program. (Not all prohibited bases apply to all 

programs.) Persons with disabilities who require alternative means for communication of program information 

(Braille, large print, audiotape, etc.) should contact 

USDA’s TARGET Center at (202) 720-2600 (voice and 

TDD). To file a complaint of discrimination, write USDA, 

Director, Office of Civil Rights, 1400 Independence 

Avenue, S.W., Washington, D.C. 20250-9410, or call 

(800) 795-3272 (voice) or (202) 720-6382 (TDD). USDA 

is an equal opportunity provider and employer.

Nomenclature for scientific names follows the U.S. Department of Agriculture, Natural Resources Conservation Service 

PLANTS (Plant List of Accepted Nomenclature, Taxonomy, 

and Symbols) database (2008). http://plants.usda.gov.

iv

Cover photo credits (left to right, top to bottom): Brian F. Daley, Jill 

Wagner, Brian F. Daley, Thomas D. Landis, George Hernández, Jill 

Wagner, Diane L. Haase, Brian F. Daley, and Thomas D. Landis. 

Back cover photo credits (top to bottom): Brian F. Daley, © Jack 

Jeffrey Photography, Tara Luna, Diane L. Haase, and J.B. Friday.
CONTENTS

Getting Started

1. Why Start a Tropical Nursery for Native and Traditional Plants?. . . . . . . . . . . . . . . . . . . . . . . . . . 17

Kim M. Wilkinson and Brian F. Daley

The Environmental and Social Context in the U.S. Affiliated Tropical Islands. . . . . . . . . . . . . . . . . . . . . . 19

Why Grow Native Plants?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20

Why Grow Traditional Plants?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

The Role of Nurseries in Ecosystem Restoration and Cultural Renewal. . . . . . . . . . . . . . . . . . . . . . . . . . . . 25

Challenge and Opportunity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

2. Planning a Tropical Nursery. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31

Kim M. Wilkinson and Thomas D. Landis

The Importance of Planning. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32

Defining the Nursery’s Vision and Objectives. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33

Determining Community Needs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33

Defining Target Plant Needs and Other Services. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35

Assessing Resources and Costs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36

Selecting a Nursery Site. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

Planning the Flow of Work. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .42

Planning as an Ongoing Process. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43

3. Defining the Target Plant . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45

Thomas D. Landis and Kim M. Wilkinson

Important Approaches for Implementing the Target Plant Concept. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46

1. What Are the Outplanting Objectives?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48

2. What Are the Conditions of the Outplanting Site?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51

3. What Factors on the Project Site Could Limit Success?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52

4. How Will Limiting Factors Be Mitigated?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53

5. What Species and Genetic Sources Will Meet Project Objectives?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55

6. What Types of Plant Materials (Stocktypes) Are Best Suited to the Project Site and Objectives?. . . . . 58

7. What Are the Best Outplanting Tools and Techniques?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63

8. What Is the Best Time for Outplanting?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63

Learning and Adapting: Field Testing the Target Plant. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65

4. Crop Planning: Propagation Protocols, Schedules, and Records. . . . . . . . . . . . . . . . . . . . . . . . . . 67

Kim M. Wilkinson and Douglass F. Jacobs

Developing Propagation Protocols. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69

Developing Growing Schedules. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72

Propagule Collection, Processing, and Treatment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72

Crop Growth Phases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73

v
Developing Facilities Schedules. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75

Keeping Written Records Makes Planning Easier. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

Appendixes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84

Developing Your Nursery

5. Propagation Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

Douglass F. Jacobs, Thomas D. Landis, Tara Luna, and Diane L. Haase

Matching Propagation Environments to the Site . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90

The Challenge of Growing Many Species and Stocktypes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90

Minimally Controlled Propagation Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

Semicontrolled Propagation Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 92

Fully Controlled Propagation Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93

Specialized Rooting Propagation Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95

Modifying Light and Temperature in Propagation Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97

Equipment Maintenance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99

Additional Reading . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99

6. Growing Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101

Thomas D. Landis, Douglass F. Jacobs, Kim M. Wilkinson, and Tara Luna

Functions of Growing Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102

Physical Properties of Growing Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103

Chemical Properties of Growing Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103

Biological Properties of Growing Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105

Growing Media Ingredients . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105

Developing a Growing Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112

Treatment of Growing Media Ingredients . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115

Mixing Growing Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117

Safety Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118

Testing Growing Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120

Additional Reading . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121

7. Containers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123

Thomas D. Landis, Tara Luna, and R. Kasten Dumroese

Container Characteristics Affecting Plant Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124

Economic and Operational Factors Affecting Container Choice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128

Types of Containers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129

vi

CONTENTS (continued)

Tropical Nursery Manual
vii

CONTENTS (continued)

Cleaning Reusable Containers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139

Plant Propagation

8. Collecting, Processing, and Storing Seeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 141

Tara Luna and Kim M. Wilkinson

Understanding Genetic Diversity and Seed Collection Ethics. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142

Understanding Flowers and Seeds. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144

Collecting Seeds. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 148

Processing Seeds. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152

Seed Testing. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 156

Storing Seeds. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 160

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161

9. Seed Germination and Sowing Options . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163

Tara Luna, Kim M. Wilkinson, and R. Kasten Dumroese

Seed Characteristics. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164

Treatments To Overcome Seed Dormancy and Enhance Germination. . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

Environmental Factors Influencing Germination. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171

Seed Sowing Methods. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

Seed Coverings (Mulch). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178

Try Different Sowing Techniques and Keep Detailed Records. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181

Larger Role of Nurseries in Conservation of Species. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183

10. Vegetative Propagation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185

Tara Luna and Diane L. Haase

Comparing Vegetative to Seed Propagation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186

Cuttings. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186

Selecting Cuttings From Mother Plants. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189

Collecting, Transporting, and Storing Cuttings. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

Types of Rooting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

Cutting Preparation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192

Striking, Monitoring, and Growing Cuttings. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194

Other Methods of Vegetative Propagation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205

Table of Contents
Plant Care

11. Water Quality and Irrigation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207

Thomas D. Landis and Kim M. Wilkinson

Sources of Irrigation Water. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208

Water Quality. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209

Water Treatments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212

Water Quantity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213

Factors Affecting Water Availability to Plants. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214

Determining How Much to Irrigate. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214

Fertigation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217

Determining When To Irrigate. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217

Types of Irrigation Systems. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220

Water Conservation and Managing Nursery Wastewater. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 230

12.Plant Nutrition and Fertilization. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233

Douglass F. Jacobs and Thomas D. Landis

Basic Principles of Plant Nutrition. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234

Sources of Mineral Nutrients. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236

Organic Fertilizers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

Synthetic Fertilizers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 240

Comparison of Organic Versus Synthetic Fertilizers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 243

Fertilizer Application Rates and Methods. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244

Determining When to Fertilize. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246

Monitoring and Testing. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248

Reducing the Environmental Effects of Fertilization. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 250

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 250

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 250

13. Beneficial Microorganisms. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253

Kim M. Wilkinson and David P. Janos

The Importance of Beneficial Microorganisms in the Nursery. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255

Nitrogen-Fixing Bacteria. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256

Mycorrhizal Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262

Other Beneficial Microorganisms. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

 Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271

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CONTENTS (continued)
14. Problem Prevention and Holistic Pest Management . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273

Thomas D. Landis, Tara Luna, R. Kasten Dumroese, and Kim M. Wilkinson

Nursery Diseases and Pests. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275

Problem Prevention Through Cultural Measures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276

Problem Detection and Diagnosis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279

Problem Management. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282

Ongoing Process Evaluation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289

Acknowledgment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291

Hardening, Shipping, and Outplanting

15. Hardening. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293

Douglass F. Jacobs, Thomas D. Landis, and Kim M. Wilkinson

The Importance of Hardening. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 294

Objectives of the Hardening Phase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 295

Conditioning Plants for Outplanting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301

16. Harvesting and Shipping. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303

Diane L. Haase, Thomas D. Landis, and Tara Luna

Scheduling the Harvesting Window. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 304

Grading Plants for Outplanting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 304

Short-Term Storage. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305

Packing. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 306

Shipping. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 306

Handling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 308

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311

17. Outplanting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313

Diane L. Haase, Thomas D. Landis, and R. Kasten Dumroese

Review of the Target Plant Concept and Implications for Outplanting. . . . . . . . . . . . . . . . . . . . . . . . . . . . 314

Site Preparation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316

Onsite Inspection, Storage, and Handling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 319

Selecting Planting Spots. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320

Proper Planting Technique. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322

Treatments at the Time of Planting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 328

Monitoring Planting Quality. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331

Monitoring Outplanting Performance. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331

Post-Planting Maintenance. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333

Long-Term Follow-up: Refining Target Plant Specifications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 335

Table of Contents ix

CONTENTS (continued)
Relationships, Management, and Learning

18. Working With People. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339

Kim M. Wilkinson

Interact Based on the Nursery Vision and Objectives. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340

Working With Yourself. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340

Working With Nursery Staff. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341

Working With Clients. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 342

Working With the Community. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 348

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349

19. Nursery Management. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351

Thomas D. Landis and Kim M. Wilkinson

Identifying Nursery Tasks. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 352

Planning and Scheduling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 352

Daily Tasks and Observations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354

Recordkeeping. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 355

Crop Planning and Production Tasks. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 356

Seasonal Cleanup and Maintenance. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357

Financial Management. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357

Problem Solving. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359

20. Discovering Ways to Improve Nursery Practices and Plant Quality. . . . . . . . . . . . . . . . . . . . . . 361

Kim M. Wilkinson and Diane L. Haase

The Importance of Trying New Things in the Nursery. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362

Making Observations and Keeping Records. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363

Step 1: Developing Ideas for Nursery Experiments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364

Step 2: Gathering Information. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

Step 3: Specifying a Question and Creating the Hypothesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

Step 4: Designing the Experiment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

Step 5: Conducting the Experiment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 368

Step 6: Making Observations and Collecting Data. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 368

Step 7: Assessing, Recording, and Sharing Results. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371

Requesting Assistance When Needed. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371

Field Testing to Refine the Target Plant. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373

Beyond Experiments: Learning Through Reflective Practice. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374

References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376

Additional Reading. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376

x Tropical Nursery Manual

CONTENTS (continued)
Brian F. Daley is a forest ecologist in the U.S. Virgin Islands. 

He was the Agroforestry Research Specialist for the University 

of the Virgin Islands for 10 years. While he was at the university, 

he developed native tree propagation protocols and low-cost 

“gap-planting” methods using native plants to enrich degraded 

dry forests dominated by exotic plants. His research includes 

landcover change analysis of St. Croix using Landsat and focusing 

on the relationship between forests, agriculture, and development. He is currently a senior partner with Geographic Consulting, 

LLC, a natural resources management company with projects ranging from phytoremediation of contaminated industrial sites to sea 

turtle nesting research and writing legislation.

R. Kasten Dumroese is the U.S. Department of Agriculture 

(USDA), Forest Service National Nursery Specialist and a research 

plant physiologist in the Rocky Mountain Research Station. His 

background as a nursery manager and researcher shapes his philosophy that nursery production problems are simply “unsolved 

opportunities.” His research focuses on collecting, growing, and 

outplanting native plants; removing bottlenecks to efficient plant 

production and quality; and correctly using native plants in 

response to climate change and to restore ecosystem function. In 

2000, he initiated the Native Plants Journal and the Native Plant 

Network; this journal and searchable Internet database foster 

sharing of information about growing and planting native flora.

Diane L. Haase is the USDA Forest Service Western Nursery 

Specialist, providing technical expertise to nurseries in the 17 

Western States and the Pacific islands. Through publications, 

presentations, conferences, workshops, and on-site visits, 

Diane assists nursery managers and other plant professionals 

in improving nursery growing practices, increasing seedling 

quality, and maximizing plant growth and survival after outplanting. She has published numerous scientific articles and 

technical papers and is a co-author of the book Propagation 

of Pacific Northwest Native Plants. In addition, she is the Editor for Tree Planters’ Notes, a journal focused on nursery production and outplanting of trees, shrubs, and native plants for 

reforestation, conservation, and restoration.

Douglass F. Jacobs is the Fred M. van Eck Professor of Forest 

Biology in the Department of Forestry and Natural Resources at 

Purdue University. He is the Editor-in-Chief of New Forests, an 

international journal on the biology, biotechnology, and management of afforestation and reforestation. His research explores 

eco-physiological development of young trees in response to 

culture and environmental stresses. He teaches conceptual 

thinking and problemsolving skills relevant to the multifaceted, 

fluid decisionmaking process of operational management for 

nurseries and land managers. Having worked in Hawai‘i for more 

than 10 years, he was closely involved in the formation of the Tropical Hardwood Tree Improvement and Regeneration Center.

David P. Janos is Professor and Cooper Fellow in the Department of Biology, University of Miami. His specialties are ecology, 

mycology, and tropical botany, and his research concerns the 

ecophysiology of mycorrhizae—mutualistic symbioses between 

fungi and the roots of vascular plants. The objective of his work is 

to understand the biology of mycorrhizae in natural ecosystems 

so that they can be exploited for human benefit in sustainable forest management, polycultural agriculture, and reforestation. As a 

former managing editor of the journal Mycorrhiza, he has worked 

extensively with the mycorrhizae of tropical tree seedlings in 

Costa Rica, Panama, and Australia.

Thomas D. Landis was the National Nursery Specialist for 

the USDA Forest Service; he taught a series of tropical nursery 

training sessions in Hawai‘i, American Samoa, and Micronesia.  

In 2003, he invited Wilkinson, Luna, and Jacobs to participate in 

teaching a tropical nursery workshop that was so successful that 

it became the impetus for this book.  Landis has authored many 

articles and books on nursery technology including Forest Nursery Notes and the Container Tree Nursery Manual series, which 

helped promote key nursery concepts including the target seedling and propagation protocols.  After retiring, he has continued 

to work as a consultant specializing in writing and training projects concerning nurseries, reforestation, and restoration.

Tara Luna is a botanist and ecologist involved in the conservation and restoration of ecologically significant areas. She 

specializes in documenting plant species and plant community 

diversity on indigenous lands, including helping to locate, protect, 

and restore areas with rare and culturally significant species. She 

has worked with American Indian tribes in the Western United 

States and has also taught native plant nursery short courses in 

the tropical Pacific. She edited Nursery Manual for Native Plants: 

A Guide for Tribal Nurseries and has authored native plant propagation, conservation, and restoration publications for the Native 

Plants Journal, USDA Forest Service and Foreign Agriculture Service, and Montana Natural Heritage Program. 

Kim M. Wilkinson is a social ecologist who works with 

ecological restoration and cultural renewal, helping bring back 

native and traditional plants, as well as human connections to 

them. She worked for 10 years in Hawai‘i as the owner/operator 

of a nursery for native and traditional trees. She also focused 

on applying traditional Polynesian agroforestry practices to 

sustainable farm and forestry planning. She continues to work 

at the intersections of food security, ecosystem restoration, and 

human health. She has authored several books, including Agroforestry Guides for Pacific Islands; Growing Koa: A Hawaiian Legacy 

Tree; and Other Voices, Other Ways, Better Practices: Bridging 

Local & Professional Environmental Knowledge. 

xi

About the Authors
xii

Acknowledgments

The U.S. Department of Agriculture (USDA), Forest 

Service funded this project. Primary funding was provided by State and Private Forestry, Cooperative Forestry, 

through the Virtual Center for Reforestation, Nurseries, 

and Genetics Resources (RNGR). The Rocky Mountain 

Research Station also supported this effort. 

Jim Marin provided graphic design and layout with 

unfailing skill, patience, and attention to detail. Richard 

Zabel, Western Forestry and Conservation Association, 

through an agreement with the RNGR team, was instrumental in developing this manual. 

J.B. Friday, College of Tropical Agriculture and Human 

Resources, University of Hawai‘i generously shared his 

expertise, insights, and many excellent photographs. 

Jill Wagner of Future Forests Nursery, Hawai‘i, contributed photographs and her expertise on nursery management and seed saving. Craig Elevitch of Agroforestry Net, 

Hawai‘i, provided photographs and his expertise in traditional trees, agroforestry, and nursery development. Ray D. 

Rodríguez Colón, Fideicomiso de Conservación de Puerto 

Rico, graciously hosted visits to his country’s nurseries and 

gave insightful input. Dania Rivera Ocasio, University of 

Puerto Rico, facilitated nursery visits. Michael Morgan of 

the University of the Virgin Islands made many thoughtful 

contributions. Vital review, fact-checking, or information 

was provided by: Robert L. James, USDA Forest Service 

(retired); Mike Amaranthus, Mycorrhizal Applications, 

Inc.; Jim Trappe, Oregon State University; RNGR team 

members George Hernández, Ron Overton, and Jeremiah 

R. Pinto; and Margie V. Cushing Falanruw, Institute of 

Pacific Islands Forestry, Yap Institute of Natural Science. 

Krystal Beley provided consultation on photograph use 

and caption clarity. We thank Katie Friday, USDA Forest 

Service Institute of Pacific Islands Forestry for contributing 

photographs and information and assisting with the tropical nursery training session that first brought our authoring team together.

We are grateful to the many individuals across the 

tropical nursery community and other professionals who 

helped create this publication: American Samoa: Aitasi 

Sameli, Endangered Plant Program; Forestry staff, American Samoa Community College; Guam: David Limtiaco, 

USDA Forest Service; Guam Forestry staff; Hawai‘i: Rick 

Barboza, Hui Kū Maoli Ola Native Plant Nursery; Mike 

Donoho, Pu‘u Wa‘awa‘a Forest Reserve; John L. Edson, 

Hawai‘i Reforestation Nursery; Jack Jeffrey Photography; 

Baron Horiuchi and his nursery staff, U.S. Department 

of the Interior (DOI), U.S. Fish & Wildlife Service Hakalau Forest National Wildlife Refuge; Joy Hosokawa, DOI 

National Park Service Volcanoes National Park; Mark and 

K.B. Kimball, Holualoa Tree Farm; Elliott Parsons, Pu‘u 

Wa‘awa‘a Forest Reserve; Ethan Romanchak, Native Nursery; Matt Schriman, Hui Kū Maoli Ola Native Plant Nursery; Jacob Witcraft and his nursery staff, Kamuela State 

Tree Nursery; Aileen Yeh, Po Wai U Nursery; Jim Ferrell, 

Mitiku Habte, Harold Keyser, and James Leary (University 

of Hawai’i); Patty Moriyasu (Volcano Rare Plant Facility); 

Idaho: J. Chris Hoag, USDA Natural Resources Conservation Service; Mississippi: J.A. Vozzo, USDA Forest Service 

Southern Research Station (retired); New York: Kenneth 

Mudge, Cornell University; Oregon: Bruce McDonald and 

Timber Press, Inc.; David Steinfeld, USDA Forest Service 

Pacific Northwest Region; Palau: Palau Forestry staff; 

Puerto Rico: Alberto Areces, Parque Dona Ines; Christian Torres Santana, USDA Forest Service International 

Institute of Tropical Forestry; U.S. Virgin Islands: Jovan 

Augustin, Geographic Consulting, Errol Chichester, Virgin Islands Department of Agriculture; Christina Gasperi, 

Art Farm; Veronica Gordon, Virgin Islands Department 

of Agriculture; David Hamada, St. George Village Botanical Garden; Cynthia Holmes, Cruzan Gardens; Dean Yanez 

(Geographic Consulting, LLC); Vanessa Forbes, Dexter 

Hipolit, and Paulino “Papo” Perez (University of the Virgin Islands); Yap: Staffs at Yap Forestry, USDA Forest 

Service Institute of Pacific Islands Forestry, Yap Institute 

of Natural Science. We also thank the many individuals 

who were photographed working in the nursery, planting 

in the field, or attending a training session.

We are grateful to the teachers, mentors, and plant 

propagators from earlier times, whose names we may not 

know but who protected and passed down knowledge of 

the native and traditional plants we enjoy today. 
xiii

Introduction

Who Is This Handbook For?

This handbook was written for anyone endeavoring 

to start and operate a nursery for native and traditional 

plants in the tropics. Because the tropics cover a vast area 

of the world, however, the scope of the handbook is geared 

toward readers in the U.S. affiliated tropics. Specifically, 

the U.S. affiliated tropics are a diverse area spanning two 

oceans and half the globe, including the nations of the Federated States of Micronesia, the Republic of Palau, and the 

Republic of the Marshall Islands, as well as the Territory 

of Guam, the Commonwealth of the Northern Mariana 

Islands, the Territory of American Samoa, the Commonwealth of Puerto Rico, the U.S. Virgin Islands, and the State 

of Hawai‘i, southern California, Texas, and the southern 

part of Florida (see map on following page). Areas with 

similar conditions may also be served. This handbook is 

not a species-by-species account, though propagation protocols for many tropical species are available on the Native 

Plant Network (http://www.nativeplantnetwork.org). 

The chapters in this handbook discuss aspects of nursery management, providing an overview of the factors 

to consider when planning or upgrading a nursery. Key 

concepts and processes are presented, based on proven 

techniques, practices, and the best science available at the 

time of this writing. An understanding of some of these 

concepts and principles will make it easier to operate a 

nursery successfully, to serve clients, and to meet project 

objectives in the field. 

At the same time, every nursery is unique. Local conditions and ingenuity, combined with these basic principles, 

will shape the best practices for any given nursery. Information in this handbook is meant to provide an empowering overview of concepts and principles. Each reader is free 

to adapt, adopt, or disregard this information, as appropriate to their situation. 

What’s in This Handbook?

A nursery is a web of interrelated factors, and every 

aspect of the nursery will affect everything else. Therefore, 

this handbook covers topics from initial nursery founding 

and planning through crop production, outplanting, and 

ongoing learning. The important roles tropical nurseries 

play in ecological restoration and human well-being (Chapter 1) are discussed before going into the details of nursery 

site selection and planning (Chapter 2). Then, the focus 

shifts to what species and stocktypes the nursery will provide to best serve different projects and objectives (Chapter 

3). The practicalities of planning crops, scheduling production activities, and developing ways to grow new crops is 

discussed (Chapter 4). Crop growth relies on good design 

of propagation environments (Chapter 5) and other infrastructure, including choices of growing media (Chapter 6) 

and appropriate containers (Chapter 7). Plant propagation 

includes collecting, processing, and storing seeds (Chapter 

8), germinating and sowing seeds (Chapter 9), and vegetative propagation (Chapter 10). Plant care includes water 

quality and irrigation management (Chapter 11), plant 

nutrition (Chapter 12), working with beneficial organisms 

(Chapter 13), and holistic prevention and management of 

pests and diseases (Chapter 14). As crops get closer to being 

ready for the field, nursery management continues with 

hardening (Chapter 15), care in harvesting and shipping of 

nursery crops (Chapter 16), and finally, outplanting onto 

field sites and monitoring the results (Chapter 17). 

All these activities combine to produce the best quality plants with the best chance to survive and flourish on 

outplanting sites. It is important, however, not to become 

so focused on production that other key aspects of nursery planning and management are neglected. Equally 

important is time building relationships with customers, the community, nursery staff, and other colleagues. 

These activities include training and education for nursery 

staff and outreach and education to clients and the public 

(Chapter 18). Overseeing nursery operations pulls many of 

the previous topics together as the nursery gains momentum in its operations (Chapter 19). Running a nursery is 

a continuous learning process and in-house research, trials, and feedback systems help improve plant productivity 

to expand on successes and learn from failures over time 

The goal of this handbook is to provide 

practical, user-friendly, science-based, and 

locally adaptable information about how to 

start, operate, or upgrade a nursery for native 

and traditional plants in the tropics.
(Chapter 20). Best efforts in each of these areas of tropical 

nursery management serve ecological, cultural, and economic goals to protect and restore native and traditional 

ecosystems and agroecosystems. 

We hope you find this handbook to be a useful reference for 

your nursery and restoration work in the tropics. 

This handbook emphasizes principles and 

processes for successful nursery operations 

that can be adapted as appropriate to 

each nursery’s context. 

! ! ! IneIne

Anekirea

Ajiltake

Ajaltokrok

15 5° 0'0 " E

15 5° 0'0 " E

16 0° 0'0 " E

16 0° 0'0 " E

16 5° 0'0 " E

16 5° 0'0 " E

17 0° 0'0 " E

17 0° 0'0 " E

17 5° 0'0 " E

17 5° 0'0 " E

6°0'0"N 6°0'0"N

11°0'0 "N 11°0'0 "N

640

Miles

65°0'0 "W

65°0'0 "W

64°3 0'0 "W

64°3 0'0 "W

18°0'0 "N 18°0'0 "N

 

St. Croix

St. Thomas

St. John

25

Miles

67°0'0 "W

67°0'0 "W

66°0'0 "W

66°0'0 "W

18°0'0 "N 18°0'0 "N

19°0'0 "N 19°0'0 "N

 

80

Miles

! !

! !

Ofu Olosega

Aunu`u

Pago Pago

17 0° 30'0"W

17 0° 30'0"W

17 0° 0'0 "W

17 0° 0'0 "W

16 9° 30'0"W

16 9° 30'0"W

14°1 8'0 "S 14°1 8'0 "S

Tutuila Ta'u

Ofu-O losega

30

Miles

!

!

Hilo

Lih ue

16 0° 0'0 "W

16 0° 0'0 "W

15 7° 30'0"W

15 7° 30'0"W

19°0'0 "N 19°0'0 "N

21°3 0'0 "N 21°3 0'0 "N

Kauai

Oahu

Maui

Hawaii

170

M i les

 

!

13 4° 30'0" E

13 4° 30'0" E

7°0'0"N 7°0'0"N

7°3 0'0 "N 7°3 0'0 "N

8°0'0"N 8°0'0"N

 20

Miles

Koror

!

!!!

!

14 0° 0'0 " E

14 0° 0'0 " E

15 0° 0'0 " E

15 0° 0'0 " E

16 0° 0'0 " E

16 0° 0'0 " E

1°0'0"N 1°0'0"N

11°0'0 "N 11°0'0 "N

 

Kosr ae

Pohnpei

Chuuk

Yap

660

Miles

Kolonia

Utwa M a

!

!

! 14 5° 30'0" E

14 5° 30'0" E

14°1 2'0 "N 14°1 2'0 "N

14°4 2'0 "N 14°4 2'0 "N

15°1 2'0 "N 15°1 2'0 "N

Rota

Tinian

Saipan

 

 

25

Miles

C ha lan K an o a

Sa n Jos e

Songso ng

!

!

!

^_

!

!

14 4° 36'0" E

14 4° 36'0" E

14 4° 54'0" E

14 4° 54'0" E

13°2 7'0 "N 13°2 7'0 "N

10

Miles

H ag at na

T amunin g

Yig o

Me rizo

Ag at Tropical Island Associates

Commonwealth

of Puerto Rico

US Virgin Islands

American Samoa

Federated States Republic of the Marshall Islands

of Micronesia

Commonwealth of the

Northern Mariana islands

Guam

State of Hawaii

Republic

of Palau

2 -40 ̊F through -50 ̊F

3 -30 ̊F through -40 ̊F

4 -20 ̊F through -30 ̊F

5 -10 ̊F through -20 ̊F

6 0 ̊F through -10 ̊F

7 10 ̊F through 0 ̊F

8 20 ̊F through 10 ̊F

9 30 ̊F through 20 ̊F

Tropical

10 40 ̊F through 30 ̊F

Zone Average Anual Low

Tropical Forests of the United States and Affiliated Islands 

Adapted from USDA Forest Service, State and Private Forestry (2007) and Arbor Day Foundation (2006).

xiv
References

Arbor Day Foundation. 2006. Hardiness zone map. http://www.arborday.org/media/zones.cfm. (October 2011).

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

U.S. Department of Agriculture, Forest Service, State and Private Forestry, Program Redesign Committee. 2007. Tropical forests of 

the United States: applying U.S. Department of Agriculture, State and Private Forestry programs. http://www.hawaiistateassessment.

info/library/tropicalforests-of-the-United-States-Final60607.pdf. (November 2011).

xv

The development of most nursery crops can be divided into three phases: establishment, rapid growth, and hardening. Adapted 

from Dumroese and others (2008) by Jim Marin.
16
17

Why Start a Tropical Nursery 

for Native and Traditional Plants?

Kim M. Wilkinson and Brian F. Daley

1

Tropical ecosystems and agroecosystems are vital, life-giving landscapes and 

are home to diverse plants, animals, and people in a range of climatic, geologic, cultural, and environmental contexts. These systems provide services, such as cleaning 

air, improving water quality, stabilizing soil, and regulating the climate. The landscapes offer essential products including timber, food, fibers, and medicines. 

During the past few centuries, waves of change have swept through tropical 

landscapes. These changes include obliterated native forests, drained wetlands, 

and decimated agroforests and indigenous management practices. Agricultural 

and industrial land uses, tourism, urbanization, climate change, and the introduction of problematic new plants, animals, pests, and pathogens have caused 

high rates of extinction (Carter and others 2001). Today, people are working to 

protect and regenerate tropical ecosystems and agroecosystems (Robotham and 

others 2004). Key tasks in revitalizing these systems are planting, protecting, and 

perpetuating native and traditional species, and sowing awareness of these plants 

among people. This is the work of the tropical nursery manager.

Nurseries in the tropics range in size from a single owner-operator growing 

a few plants for neighbors, to community organizations, universities, or cultural 

groups growing plants for certain goals, to large commercial nurseries, to governments founding native plant programs for their region. Without regard to scale, 

tropical nurseries are a key link in ecological restoration, sustainable agriculture, 

and cultural renewal (figure 1.1) by providing native and traditional plants.

Facing Page: Nursery on Pohnpei, Federated States of Micronesia. 

Photo by Ronald Overton.
18 Tropical Nursery Manual

diverse native and traditional species for timber, crafts, 

fibers, foods, and other uses; diversifying monoculture 

fields with traditional species of trees, shrubs, and vines; 

using native and traditional plants in conservation 

plantings, such as windbreaks, urban street trees, and 

erosion control installations.

• Learning and Cultural Renewal—Rekindling locally 

adapted practices and plant technologies; cultivating 

culturally significant plants in the forest understory 

for nontimber forest products; replacing exotic 

ornamentals with local species in landscapes; starting gardens of traditional species in schoolyards; 

revitalizing and sustaining human connections with 

local plants and ecosystems (figure 1.2).

Examples of these links include—

• Ecosystem Restoration and Biodiversity Protection—

Jump-starting or accelerating succession on disturbed 

sites; enriching species-poor secondary forests with rare 

or later successional species; partnering with farmers 

and agroforestry practitioners to facilitate tropical forest 

recovery; bringing native plants for native pollinators 

and wildlife into agricultural areas, urban landscapes, 

and backyard gardens; propagating threatened and 

endangered plants for their conservation and reintroduction to forests; and the full-scale restoration of native 

plant communities in conservation areas.

• Sustainable Agriculture and Economic Resilience—

Enhancing farmer livelihoods and community food 

security with agroforestry practices; rediscovering 

Figure 1.1—Nurseries in the tropics come in many shapes and 

sizes to serve diverse needs. Shown here are native tree seedlings 

in a nursery in Guam (A), a backyard nursery in Mexico (B), and 

a commercial nursery in Puerto Rico (C). Photo A by J.B. Friday, 

photo B by Thomas D. Landis, and photo C by Ronald Overton. 

Figure 1.2—Nurseries play a key role in ecological restoration, cultural renewal, economic resilience, and sustainable agriculture 

by providing native and traditional plants. Nursery-grown plants are often used for planting native species in the U.S. Virgin Islands 

(A) and Hawai‘i (B). Photo A by Brian F. Daley, and photo B by J.B. Friday.

B

B

C

A

A
Why Start a Tropical Nursery for Native and Traditional Plants? 19

The Environmental and Social 

Context in the U.S. Affiliated 

Tropical Islands

One of the greatest challenges of nursery management is 

choosing how to focus your efforts to be of most benefit to 

the land and the local people. Understanding your context 

requires integrating a “bird’s-eye view” and a “worm’s-eye 

view.”A quick “bird’s-eye view” of the context in the U.S. 

Affiliated Tropical Islands is shown in table 1.1; you are 

the only one who can describe your unique local context 

(the “worm’s-eye view”).

The U.S. Affiliated Tropical Islands house an immense 

diversity of native ecosystems, including mangrove swamps, 

wetlands, riparian areas, rainforests, dry forests, submontane forests, cloud forests, and more (table 1.2). These ecosystems are home to many unique, endemic plants found 

nowhere else on earth and are considered biodiversity 

hotspots of local and global significance in relatively small 

areas (figure 1.3). The health of the terrestrial areas affects 

diverse marine systems including estuaries, salt ponds, and 

coral reefs. These systems provide resources for a tremendous range of marine animals and critical breeding grounds 

for migratory and residential birds. The rich and unique biological diversity of these islands is an international heritage.

Islands are particularly susceptible to environmental changes and disturbances. The small populations of 

endemic plants and animals that evolved in isolation are 

unable to change locations or rapidly adapt when large 

changes occur. Changes facing island ecosystems include 

habitat destruction because of human development 

(urban expansion, agriculture, industrial use, and recreational use) and loss of habitat because of introductions 

of exotic invasive species (plants, animals, and diseases). 

Islands are extremely vulnerable to temperature shifts, 

altered or increased storm regimes, floods, and droughts 

associated with climate change. On islands, many native 

plants and animals are endangered; the islands are in an 

extinction crisis, experiencing the highest extinction rates 

anywhere in the United States (Carter and others 2001).

Throughout the tropics, native forests were cleared and 

converted to agriculture and then abandoned, leading to 

widespread creation of secondary forests. In the Caribbean, 

where land area is limited and human population density is 

Name

Status 

with 

United 

States

Geologic 

description

Total 

area 

(acres)

Forested 

area 

(acres)

Estimated 

human 

population 

(2010)

Commonwealth of 

Puerto Rico Territory One volcanic main island composed of uplifted 

sedimentary rocks and several smaller islands 2,199,901 710,156 3,721,208

U.S. Virgin Islands Territory

Three volcanic main islands (St. Croix, St. John, and St. 

Thomas) composed of uplifted sedimentary rocks and 

several smaller islands and cays

85,587 52,477 106,267

Hawaiian Islands State

Eight volcanic islands (Kaua‘i, O‘ahu, Moloka‘i, 

Lana‘i, Maui, Kaho‘olawe, Ni‘ihau, and Hawai‘i) and 

several atolls

4,110,720 1,490,000 1,360,301

American Samoa Territory Five volcanic islands and two coral atolls (Ofu,Ta’u, 

Swains Island, Tutuila, Olosega, Rose Island, and Aunu’u) 49,280 28,686 55,467

Commonwealth of the 

Northern Mariana Islands Territory 14 volcanic islands 113,280 40,000 53,517

Republic of the 

Marshall Islands FAS* 5 low-lying volcanic islands and 29 coral atolls 

(each made up of many islets) 44,800 Not 

mapped 65,859

Republic of Palau FAS* Several hundred volcanic islands and a few 

coral atolls (eight islands inhabited) 114,560 77,241 20,879

Federated States of 

Micronesia FAS* 607 small islands consisting of volcanic islands and 

coral atolls 149,804 76,527 107,154

Guam Territory One volcanic island composed of uplifted 

sedimentary rocks 135,680 65,005 159,434

Table 1.1—Characteristics of U.S.-Affiliated Tropical Islands. Adapted from USDA FS SPF Program Redesign Committee (2007), 

Carter and others (2001), U.S. Census Bureau (2014a, 2014b), and Brandeis and Oswalt (2007). 

* Freely Associated State in a Compact of Free Association with the United States.
20 Tropical Nursery Manual

high, secondary forests on old agricultural land are already 

the most important and rapidly expanding forest cover type 

(Lugo and Helmer 2004). Secondary forests have novel 

species compositions, lower native species diversity, and 

higher influence of exotic species. In many areas in the 

Pacific Islands, secondary forests are composed exclusively of exotic species. Secondary forests will likely play 

an increasing role in conservation of forest biodiversity, 

especially in fragmented landscapes (Brown and Lugo 

1990; Chazdon and others 2009).

Inextricably linked to these diverse ecosystems are diverse 

traditional cultures of the islands. A few examples include 

the Chamorro cultures of Guam and the Mariana Islands, the 

Samoan and Hawaiian cultures, the Boricua culture of Puerto 

Rico, and cultural groups who speak multiple languages and 

call the islands of the Federated States of Micronesia home 

(Carter and others 2001). Most of the islands have significant 

populations practicing traditional lifeways, including agricultural practices, hunting, fishing, gathering, and resource 

management. Agroforestry systems are, on some islands, a 

key forest type (figure 1.4). Traditional resource management in the islands espouses a ridge-to-reef perspective, the 

wisdom of which has lessons for watershed protection anywhere on earth (USDA FS SPF 2007).

Within the past few centuries, immigration and migration have added to the cultural diversity of the islands. Local 

culture is now infused with, for example, the influence of 

the Africans, Danish, French, and Spanish in the Caribbean; 

the Asians and Americans in Hawai‘i; the Spanish in Guam; 

and countless others in other islands. Lifestyles in the islands 

range from densely populated urban areas to remote villages 

on outer islands (Carter and others 2001). Many islands have 

urban areas, agriculture, forest, and residential neighborhoods in close proximity to each other. Most of the islands 

currently experience a high level of dependency on mainland 

economies for food, energy, and other resources, but many 

also maintain a strong foundation and knowledge base of traditional subsistence agriculture and resource management.

Interconnections between social and ecological conditions are more evident to islanders than they are to people 

who live on large continents. Nurseries may provide native 

plants, traditional species, and other species to serve both 

humans and nature. Nurseries operate in urban, rural, 

and village settings, often integrating goals of economic 

resiliency, ecosystem protection and restoration, cultural 

renewal, and other community and ecosystem needs.

Why Grow Native Plants?

Native plants are the backbone of ecosystem function 

and integrity from soil nutrient cycling, clean water, and 

carbon sequestration to higher wildlife interactions. Individual species have evolved during the millennia to the 

Table 1.2—Some ecosystems in the U.S.-Affiliated Tropical Islands, present or historically present. Adapted from US OTA (1987).

Federated States of Micronesia

Puerto 

Rico

U.S. Virgin 

Islands

American 

Samoa Hawai‘i* Guam Northern 

Marianas Marshalls Palau Kosrae Pohnpei Yap Truk

Cloud forest 4 4 4 4 4 4 4 4

Submontane rainforest 4 4 4 4 4 4

Lowland rainforest 4 4 4 4 4 4 4 4 4 4

Riverine and 

swamp forest 4 4 4 4 4 4 4 4 4 4

Subtropical moist/

seasonal forest 4 4 4

Subtropical dry forest 4 4 4

Scrub 4 4 4 4 4 4 4 4 4 4 4 4

Wetlands 4 4 4 4 4 4 4 4 4 4 4 4

Mangrove forest 4 4 4 4 4 4 4 4 4 4 4

Atoll/beach forest 

and scrub 4 4 4 4 4 4 4 4 4 4 4 4

Lagoons/shallow bottoms 4 4 4 4 4 4 4 4 4 4 4 4

* Hawai‘i also has subalpine and alpine ecosystems.
Why Start a Tropical Nursery for Native and Traditional Plants? 21

Figure 1.3—The U.S.-Affiliated Tropical Islands house an immense diversity of ecosystems including mangrove swamps, wetlands, 

rainforests, dry forests, submontane forests, cloud forests, and more: mixed native and introduced forest on O‘ahu (A), dryland forest 

on Hawai‘i (B), coastal pandanus (screw pine) forest in Hawai‘i (C), and Rock Islands of Palau (D). Photos A, B, and C by J.B. Friday, 

and photo D by Thomas D. Landis.

Figure 1.4—Diverse agroecosystems such as agroforests are important to environmental health and human livelihood in the tropics. Snow on Mauna Kea (14,000-ft [4,600-m] elevation) framed by palm trees and breadfruit growing at 700-ft (215-m) elevation 

on the Big Island of Hawai‘i (A); an agroforest in Palau with betel nut palm (Areca catechu) overstory, bamboo for building materials, bananas and avocados for fruit, and taro for starch (B); mixed land use in the U.S. Virgin Island with residential, pasture, 

forest, and commercial areas in close proximity (C); and Coconut Island in Hawai‘i (D). Most visitors to the tropics do not realize 

that many of the landscapes they enjoy include plant species humans introduced and cultivated for centuries in the area. Photos A, 

B, and D by J.B. Friday, and photo C by Brian F. Daley.

B

B

C

C

A

A

D

D
climatic and edaphic conditions where they now grow in 

association with a myriad of other species. The resulting 

assemblage of plants and animals is what makes each ecosystem unique and contributes to people’s sense of place. 

When species associations are disrupted, local extinctions 

can occur (figure 1.5), leading to declining ecosystem 

function or even to ecosystem collapse (Hawkins and others 2008, USDI BLM SOS 2011). Restoring durable, selfsustaining, and resilient ecosystems depends on native 

plants to (Erickson 2008, 2010)—

• Recover natural vegetative composition, structure, 

and successional patterns.

• Protect biodiversity and plant genetic resources.

• Provide habitat connectivity.

• Maintain plant and pollinator interactions.

• Support wildlife populations.

• Increase system resilience to disturbances and stressors.

• Provide desired goods, services, and benefits.

• Provide economic benefit to rural communities.

In this handbook, native plants are defined as locally 

adapted, genetically appropriate native plant materials. 

Although that sounds simple, it is not a straightforward 

definition (see next section). The issue of plant nativity is 

hotly discussed in some areas; and this handbook does not 

intend to solve these debates, only to provide a working definition. In general, making a list of native plants for a region 

or an island may not be sufficient information because it 

does not indicate local adaptedness. Many widely distributed native species have local populations adapted to local 

conditions including climate, soils, elevation, precipitation, 

and environmental stresses, such as wind or drought. Local 

native plant materials, collected from the same or similar 

habitats as the outplanting site, have been shown to perform 

better than nonlocal sources. Using locally adapted seed 

sources is a key factor in ensuring the survival of native 

plants and of the native fauna that depend on these plants. 

In addition, deploying a diversity of locally adapted native 

plants creates communities and ecosystems that are more 

adaptable and resilient to climate change (Horning 2011, 

Erickson and others 2012). For some species, seed zones or 

transfer guidelines may have been defined (as discussed in 

Chapter 3, Defining the Target Plant), but establishment 

of seed zones is rare for most native tropical species. For 

most projects, growers must decide on a case-by-case basis 

what will be appropriate for their outplanting sites (Withrow-Robinson and Johnson 2006). The genetics guidelines 

provided in Chapter 8, Collecting, Processing, and Storing 

Seeds, detail essential elements of seed collection to ensure 

local adaptedness and genetic diversity.

Why Grow Traditional Plants?

This handbook recognizes that traditional plants are a 

sustainable and regenerative resource in the tropics, and 

that tropical nurseries work to serve those needs in addition to native plant conservation. Many people define 

“native” as those species present before any humans 

inhabited an area, but humans have resided in many 

parts of the tropics for millennia and have influenced 

the flora. Research is revealing that many tropical forests 

once thought to be undisturbed primary forests (such as 

in the Brazilian Amazon) are actually the result of ongoing human management and disturbances dating back, 

in some cases, for thousands of years (Berkes 1999) The 

first people to migrate to a new area brought many plants 

with them.

For example, in the Caribbean, Amerindians with 

cultural centers in Venezuela traveled by small boats 

northward, bringing with them important food crops, 

including trees such as the genip (Melicoccus bijugatis) 

and zapote (Manilkara zapota), which were then cherished for their fruit, much as they are today. Both species 

are now naturalized throughout the Caribbean Islands. 

When the first Polynesians arrived to the Hawaiian 

Islands, they found few edible native plants. The abundant agroecosystems of breadfruit, coconut, bananas, 

taro and other species that early Europeans thought to 

be native were in fact intentionally introduced species in 

Polynesian agroforestry systems. Islanders of the Pacific 

and Caribbean enjoyed a high level of food security and 

self-sufficiency before European contact (Thaman and 

22 Tropical Nursery Manual

Figure 1.5—Deploying a diversity of locally adapted native 

plants helps support and regenerate ecosystems for local flora 

and fauna. Pictured is the ‘i‘iwi (Vestiaria coccinea) drinking 

nectar from an ‘Ōhi‘a lehua (Metrosideros polymorpha) blossom 

in Hawai‘i. Photo ©Jack Jeffrey Photography.
others 2000). Their agroecosystems were built around a 

diverse base of native and introduced species (Elevitch 

and Wilkinson 2000). Many of these agroecosystems were 

displaced by colonial and later corporate entities, to convert land for development or to export crops such as sugar, 

coffee, or cattle. Today people are working to revitalize and 

reintroduce traditional species and agroecosystems for personal health and for economic, environmental, and political 

reasons (figure 1.6). For example, the U.S. Virgin Islands 

have seen a recent resurgence in Ital cooking that follows traditional Rastafarian practices of using locally grown herbs 

and vegetables without meat. Native Hawaiians are successfully overcoming nutrition-related disorders, such as 

obesity and diabetes, by reembracing traditional foods. The 

ancestral way of eating is cultivated in an interdisciplinary 

way involving cultural teachings and family support (Shintani and others 1994). On the islands, the general trend is 

toward locally grown foods in response to higher food 

prices, the desire to eat more organically and healthfully, 

to increase household food security, and to support food 

sovereignty movements. In the latter, local people claim the 

power to determine their own food systems as a basic human 

right, rather than allowing food systems to be controlled by 

global market forces (Whittman and others 2010). Growing 

traditional species in nurseries helps strengthen and renew 

sustainable agroecosystems (figure 1.7).

Helping to meet human needs using time-tested, traditional species also reduces pressure on more intact tropical 

ecosystems, without the risk of newly introduced and potentially invasive or noxious species. Traditional species can 

Why Start a Tropical Nursery for Native and Traditional Plants? 23

Figure 1.6—Traditional plant species sustain lifeways and livelihoods in 

tropical areas, providing food security, 

building materials, medicines, crafts, 

and more: a display of crops from an 

agroforestry system in the U.S. Virgin 

Islands, including a native passion fruit 

vine, medicinal neem tree leaves (originally introduced from India), and sweet 

potatoes (A); these food plants are part 

of traditional cooking and food security 

in the U.S. Virgin Islands (B). Traditional species also provide material for 

economic livelihoods, such as weaving 

materials for this traditional basket 

weaver on Yap (C). Specific native trees 

are used for building traditional fales 

in American Samoa (D and E). Photos 

A and B by Brian F. Daley, photo C by 

Thomas D. Landis, and photos D and E 

by Diane L. Haase. 

B

E

C

A D
also be an integral part of tropical forest restoration using 

what is known as an agro-successional restoration approach 

(Vieira and others 2009, Holl and Aide 2010). In this 

approach, agroforestry and other agroecological practices 

are employed as a transition phase for forest restoration. 

Farmers share in restoration efforts to improve soil, establish 

trees, and exclude weeds while meeting their own needs for 

food security and livelihood (Vieira and others 2009). In this 

handbook terms including “traditional,” “culturally significant,” and “culturally important” are used interchangeably 

to refer to these kinds of plant species; some gray areas exist 

as to which species are in this category. Some culturally 

significant species are also native species; some are introduced species. Of the introduced species, some arrived with 

the first peoples of a place; others came with later waves 

of settlement. Some were adopted after European contact. For example, enslaved Africans arrived by ship to the 

Caribbean during the centuries of European colonization. 

Although most of the enslaved traveled without personal 

possessions, they somehow smuggled items that were culturally and spiritually important, including tree seeds. The 

native African baobab tree (Adansonia species) arrived in 

the Caribbean this way. Today in the U.S. Virgin Islands, 

the giant baobab trees (figure 1.8) serve as a tangible connection between modern Virgin Islanders and the first people who arrived from Africa and planted these tree seeds. 

A traditional species may not be of indigenous use, but 

may still be culturally significant. As a nursery manager, 

you will need to consider your scope to determine what 

species are appropriate to prioritize and perpetuate based 

on your area’s unique environmental and cultural context.

Of course it is unwise to introduce to an island any species that is harmful (invasive, noxious, or untested in the 

area). In most cases, intentionally introducing new species is also unnecessary, given the diversity of time-tested 

plants already present.

24 Tropical Nursery Manual

Figure 1.7—Growing traditional species in nurseries helps strengthen and renew sustainable agroecosystems: taro paddy in 

Waipio Valley, Hawai‘i (A), shade-grown coffee under native ‘ōhi‘a forest shade, Kona, Hawai‘i (B). Photos by J.B. Friday.

Figure 1.8—Seeds of the baobab tree (Adansonia digitata) were 

first brought to the Caribbean by Africans, who somehow managed 

to carry viable tree seeds through the ordeal of enslavement and 

travel on slave ships, and who planted their seeds upon arrival. 

Today the baobab is a culturally important tree in the Caribbean, 

linking today’s islanders with centuries of African heritage in the 

region. Photo by Brian F. Daley.

A B
The Role of Nurseries in 

Ecosystem Restoration and 

Cultural Renewal

Nurseries Provide Quality Plant 

Materials to Facilitate Sustainable 

and Regenerative Practices

The nursery environment is a place to germinate, grow, 

and protect locally adapted seedlings until they are healthy, 

strong, and large enough to meet the challenges of outplanting sites and achieve project goals (Evans 1996). The work 

nurseries do to produce high-quality native and traditional 

plants helps perpetuate these species (figure 1.9). Nurserygrown native plants overcome barriers to natural recolonization, such as weed competition, animal predation, and lack 

of a natural seed bank. Nursery-grown traditional species 

ensure heritage varieties, and culturally important characteristics are passed on to the next generation. Good nursery 

work makes successfully establishing native and traditional 

plants more effective, affordable, and more likely to happen, 

whether nursery clients are farmers, gardeners, restoration 

ecologists, community groups, or others. Quality plants can 

mean the difference between people believing it is too hard 

to plant native and traditional species, and having an attitude 

that they can do this (figure 1.10).

Nurseries Serve as a Cornerstone of 

Ecological Restoration

Without human disruption to their health, tropical ecosystems regenerate naturally after disturbances through the 

process of succession. Forests can be degraded or damaged 

by humans to the point where they require some help, however, and that is where nurseries come in. Often, the intensity, duration, or extent of the disturbance results in a lack 

of native propagules (seedlings, seed bank in the soil, and so 

on) on the site or within a distance that they will naturally 

disperse to revegetate the disturbed site (Holl and Aide 2010). 

In addition, limiting factors on the site, such as takeover by 

highly competitive weeds, low soil fertility, excessively compacted soil, or nonnative animals that eat seeds and young 

plants can arrest natural succession until these conditions are 

overcome (Vieira and others 2009). In these scenarios, the 

planting of nursery stock is a cornerstone of ecological work.

Why Start a Tropical Nursery for Native and Traditional Plants? 25

Figure 1.9—Nurseries play a key role in conservation and restoration of species and ecosystems. This Department of Natural Resources 

nursery in Puerto Rico is growing threatened and endangered species. 

Photo by Ronald Overton. 

Figure 1.10—The work 

nurseries do to produce highquality native and traditional 

plants helps break down 

barriers to planting and perpetuating these species. Good 

nursery work makes outplanting more effective and 

affordable for clients. Pictured: tree planters head out 

to plant native trees in Brazil 

(A) and Hawai‘i (B). Photos 

by Douglass F. Jacobs.

A B
Nursery plants may be used to (Holl and Aide 2010, 

Vieira and others 2009)—

• Jump-start succession (for example, planting trees 

to shade out aggressive weeds or improving soil with 

nitrogen-fixing pioneer native species).

• Stabilize highly disturbed areas and minimize further 

environmental damage, such as erosion.

• Enrich species-poor secondary forests or pioneered 

sites with rarer, more diverse, or later successional 

species.

• Serve as “islands” to act as dispersal points for propagules to adjacent land in need of restoration.

• Establish trees through an agro-successional approach.

• Complete full-scale restoration projects.

Nurseries Provide Livelihood and 

Learning Opportunities for People

Nurseries and nursery mangers are a bridge between 

the natural world and human needs. For example, perhaps 

an important cultural plant has become rare, and, because 

elders are unable to gather it, the children are not learning 

about this species and the local economy is no longer benefitting. The nursery can identify the species, learn how to 

propagate it, and help people to replant it, thus serving the 

local culture, economy, and ecology.

Some nurseries provide plants, employment, and knowledge to serve certain cultural groups, such as women, youth, 

or farmers. In many areas, nurseries tend to become centers 

and gathering places for people to connect with each other 

and learn about native and traditional species (PRAP 1999) 

(figure 1.11).

One of the most rewarding objectives for growing and 

outplanting native and traditional plants is the education of 

young people. Some native plant nurseries are operated by 

schools with a primary objective of environmental education, 

and other nurseries simply make themselves available to 

school groups. Young children can learn the names and uses 

of native and culturally important plants, and older children 

can enjoy the art and science involved in growing and outplanting different species. Gaining familiarity with species 

is a useful gateway to a deeper understanding of traditional 

ecological knowledge and resource management.

Nurseries Are a Key Link in Biodiversity 

Conservation

Loss of tropical biodiversity is a cause for serious concern. High levels of endemism in relatively small spaces 

mean that local habitat destruction can put already vulnerable species at greater risk of extinction. Nursery work to 

propagate and increase rare species is essential for conservation and reintroduction of these species (Prance 2005).

The restoration of disturbed sites is vital to conservation success, but so is the establishment of seed banks, 

including nurseries as living seed banks, to serve future 

needs. Conventional seed banks can conserve only species 

having “orthodox” seeds that can be dried and placed in 

long-term storage. Most tropical species, however, have 

26 Tropical Nursery Manual

Figure 1.11—Nurseries can be wonderful places for working and learning about plants (A) and passing on knowledge and values to 

the next generation (B). Photo A by Brian F. Daley, and photo B by Megan Parker.

A B
Why Start a Tropical Nursery for Native and Traditional Plants? 27

Time-Lapse Glimpses of the Tarzan River Savanna Area of the Cotal Reserve, Guam

Dr. Margie Cushing Falanruw recalls her ancestral knowledge that became a basis for a vegetation strategy for 

southern Guam. She writes (2002 p. 11): 

“In the 1950s and early 1960s, the savanna above and along the trail leading to upper Tarzan falls in Guam was a childfriendly wonderland of shrubs. Some, such as the weird Geniostoma micranthum and lovely shiny-leaved Decaspermum 

fruticosum, grew to heights of about 7 feet and formed a dense maze through which children and adults could walk. ‘Abas 

duendes’ Gloichidion bushes with bright red tip leaves produced little ‘guavas for elves.’ The bonsai-like Myrtella bennigseniana, and the pink-flowered Melastoma malabathricum grew lower than a child and were interspersed with lavender, 

Spathoglottis ground orchids, yellow flowered Curculigo orchioides, and blue flowered Dianella lilies. The ‘miniature iba 

tree,’ Phylannthus saffordi, was present along with the ‘out of focus wolf paw plant’ Lycopodium cernuum, which was 

used to decorate nativity scenes in those days. We made garlands of Cassytha, Lygodium, or Gleichenia vines to shield 

our heads from the sun, soothed our eyes with isotonic drops of liquid from Scaevola taccada berries, and sipped nectar 

from Stachytarpheta flowers. We learned to stop bleeding with crushed Glochidion leaves and to heal bruises and sprains 

with lada Morinda leaves warmed on a campfire. We walked through soft Dianella grass, enjoyed the scent of wild mint 

and the ‘chewing gum smell’ of methyl salicicate-laced Polygalla roots. We rested below gagu, the ‘singing tree’ Casuarina 

equisetifolia and listened to the ‘sound of silence’ as wind blew through its needle branches.

Most of these species were present in the 1970s when my flora and fauna class compiled a list of species in the area 

on July 18, 1970. By the 1980s however, the area was invaded by invasive Pennisetum grass, which may have increased 

its vulnerability to wildfires that were reported to have swept through the area. The burned area had been subsequently planted to Acacia trees. I no longer lived on Guam at that time. When I visited The Tarzan river area with my 

grandchildren on Saturday, August 27th, 2002, most of the savanna species were no longer as abundant and do not 

grow as tall nor luxuriant as they did before the area was invaded by Pennisetum and fire.

The Tarzan river area of the Cotal Reserve has refreshed five generations of my family, and I hope that it can be 

restored to its former state so that my great grandchildren and their children may also experience this remnant bit of 

old Guam.”

Using that ancestral knowledge, the restoration strategy in table 1.3 was developed.

Table 1.3—Revegetation strategy. Adapted from Bell and others (2002).

Type of site Problem Restoration/stewardship objectives

Shaded fuelbreaks

Grass dominated 

watersheds are unbroken 

expanses that burn readily and 

at high intensity.

Compartmentalize contiguous fuels to reduce fire size and provide enhanced 

control opportunities. Establish shaded fuelbreaks of Acacia species in a 

strategically designed pattern to compartmentalize fuels.

Swordgrass grasslands

Swordgrass is highly 

flammable, fires tend to 

kill other vegetation and contribute to the spread 

of swordgrass.

Convert portions of swordgrass dominated grasslands to forests. Plant and 

maintain buffers of native trees. Underplant Acacia plantations with native 

trees. Plant nursery-grown natives Callophyllum inophyllum, Intsia bijuga, 

Hibiscus tiliaceus, Neiosperma oppositifolia, and Pandanus tectorius adjacent to 

existing ravine forests.

Native savanna shrub 

and ravine forest

Erosion of biodiversity as a 

result of fire, feral animals, and 

invasive species.

Protection and restoration. Weed out invasive plants, protect from fire. 

Assist or enhance regeneration of native trees, shrubs, and plants. Nursery 

propagation of native species to assist with natural regeneration of uncommon 

and rare species.
“recalcitrant” seeds, meaning they cannot be dried and 

stored over extended periods (Kettle and others 2011). Thus, 

successful restoration and conservation of tropical forests 

requires that nurseries propagate and maintain genetically 

diverse living banks of recalcitrant species (Kettle and others 

2011, Merritt and Dixon 2011).

Growing plants in the nursery is far more efficient than 

attempting to establish seeds through direct seeding in the 

field. Nurseries make the most efficient use of limited seeds by 

ensuring that each one receives the necessary care and attention to survive and develop into a mature plant. Also, nursery 

propagation does not deplete naturally regenerating areas 

because it does not involve uprooting wildings to transplant.

Good nursery management supports genetic and sexual 

diversity to ensure conservation of plants and all the other 

life forms that depend on them, from pollinators to primates. Genetic diversity is also essential for future adaptability and resilience in the face of climate change. Protecting local biodiversity and ecosystem resilience also protects 

local cultural diversity and economic resilience.

Challenge and Opportunity

Much needs to be done to restore tropical forests, 

recharge aquifers, regenerate wetlands, revitalize local 

economies, rekindle traditional practices, reclaim wastelands, and renew human connections to nature. The challenges may seem overwhelming. It is fortunate that even 

the loftiest goals can be reached with practical, persistent, 

everyday work (figure 1.12). Starting and operating a nursery for native and traditional species is a vital, positive 

endeavor full of challenge and opportunity. Good nursery 

work contributes to species conservation, ecosystem resiliency, cultural diversity, enhanced human livelihoods, and 

greater health and productivity of the land.

References

Bell, F.; Falanruw, M.; Lawrence, B.; Limtiaco, D.; Nelson, D. 

2002. Vegetation strategy for southern Guam. Draft (September 

2002). Honolulu, HI: U.S. Department of Agriculture, Forest Service and Natural Resources Conservation Service; Government of 

Guam Division of Forestry.

Berkes, F. 1999. Sacred ecology: traditional knowledge and 

resource management. London: Taylor and Francis. 209 p.

Brandeis, T.; Oswalt, S. 2007. The status of the U.S. Virgin Island’s 

forests 2004. Resource Bulletin SRS-122. Ashville, NC, U.S. 

Department of Agriculture, Forest Service, Southern Research 

Station. 62 p.

Brown, S.; Lugo, A. 1990. Tropical secondary forests. Journal of 

Tropical Ecology. 6: 1–32.

Carter, L.M.; Shea, E.; Hamnett, M.; Anderson, C; Dolcemascolo, 

G; Guard, C; Taylor, M.; Barnston, T; He, Y; Larsen, M.; Loope, 

L.; Malone, L.; Meehl, G. 2001. Potential consequences of climate 

variability and change for the U.S.-affiliated islands of the Pacific 

and Caribbean. In: The potential consequences of climate variability and change: foundation report. Report by the National Assessment Synthesis Team for the U.S. Global Change Research Program. Cambridge, United Kingdom: Cambridge University Press: 

315–349. Chapter 11. http://www.usgcrp.gov/usgcrp/Library/

nationalassessment/11Islands.pdf. (December 2011).

Chazdon, R.L.; Peres, C.; Dent, D.; Shell, D.; Lugo, A.E.; Lamb, 

D.; Stork, N.E., Miller, S.E. 2009. The potential for species conservation in tropical secondary forests. Conservation Biology. 23: 

1406–1417.

Elevitch, C.R.; Wilkinson, K.M., eds. 2000. Agroforestry guides for 

Pacific Islands. Holualoa, HI: Permanent Agriculture Resources. 

http://www.agroforestry.net/afg/book.html. (December 2011).

Erickson, V.J. 2008. Developing native plant germplasm for 

national forests and grasslands in the Pacific Northwest. Native 

Plants Journal. 9: 255–266.

Erickson, V.J. 2010. Personal communication. Pendleton, OR. 

Geneticist/Native Plant Program Manager, U.S. Department of Agriculture, Forest Service, Pacific Northwest Region.

Erickson, V.J.; Aubry, C.; Berrang, P.; Blush, T.; Bower, A.; Crane, 

B.; DeSpain, T.; Gwaze, D.; Hamlin, J.; Horning, M.; Johnson, R.; 

Mahalovich, M.; Maldonado, M.; Sniezko, R.; St. Clair, B. 2012. 

Genetic resource management and climate change: genetic 

options for adapting national forests to climate change. Internal 

document. Washington, DC: U.S. Department of Agriculture, 

Forest Service. http://www.fs.usda.gov/Internet/FSE_DOCUMENTS/stelprdb5368468.pdf. (September 2012).

28 Tropical Nursery Manual

Figure 1.12—Laulima: “Many hands working together.” The 

newly minted sign for Hakalau Forest National Wildlife Refuge’s 

Native Plant Nursery, Hawai‘i. Photo by J.B. Friday.
Evans, J. 1996. Plantation forestry in the Tropics. Oxford, United 

Kingdom: Clarendon Press. 403 p.

Falanruw, M. 2002. Appendix 3. Time-lapse glimpses of the Tarzan River savanna area of the Cotal Reserve. In: Bell, F.; Falanruw, 

M.; Lawrence, B.; Limtiaco, D.; Nelson, D. Vegetation strategy 

for southern Guam. Draft (September 2002). Honolulu, HI. U.S. 

Department of Agriculture, Forest Service and Natural Resources 

Conservation Service; Government of Guam Division of Forestry.

Hawkins, B.; Sharrock, S.; Havens, K. 2008. Plants and climate 

change: which future? Richmond, United Kingdom: Botanic Gardens Conservation International. 96 p.

Holl, K.D.; Aide, T.M. 2010. When and where to actively restore 

ecosystems?  Forest Ecology and Management. 261(10): 1558–

1563.

Horning, M. 2011. Personal communication. Bend, OR. Plant 

Geneticist, U.S. Department of Agriculture, Forest Service, 

Pacific Northwest Research Station.

Kettle, C.J.; Burslem, D.F.R.P.; Ghazoul, J. 2011. An unorthodox 

approach to forest restoration. Science. 333: 36.

Lugo, A.; Helmer, E. 2004. Emerging forests on abandoned land: 

Puerto Rico’s new forests. Forest Ecology and Management. 190: 

145–161.

Merritt, D.J.; Dixon, K.W. 2011. An unorthodox approach to forest restoration—response. Science 333: 36–37. 

Pacific Regional Agricultural Programme [PRAP]. 1999. Pacific 

agroforestry: an information kit. Suva, Fiji Islands: Pacific 

Regional Agricultural Programme.

Prance, G. 2005. Foreword. In: Lilleeng-Rosenberger, K.E. Growing Hawai‘i’s native plants. Honolulu, HI: Mutual Publishing.

Robotham, M.P.; Mas, E.; Lawrence, J.H.; Eswaran, H. 2004. The 

Tropical Natural Resources Technology Consortium: working 

together for tropical conservation. In: Raine, S.R.; Biggs, A.J.W.; 

Menzies, N.W.; Freebairn, D.M.: Tolmie, P.E. eds. Conserving 

soil and water for society: sharing solutions. ISCO International 

Soil Conservation Organisation Conference, July 2004, Brisbane, 

Australia: ASSSI/IECA. http://www.ttc.nrcs.usda.gov/news/

TTC_paper_ISCO.pdf. (June 2011).

Shintani, T.; Beckham, S.; O’Connor, H.K.; Hughes, C.; Sato, A. 

1994. The Waianae diet program: a culturally sensitive, community-based obesity and clinical intervention program for the native 

Hawaiian population. Hawai‘i Medical Journal. 53: 136–147.

Thaman, R.R.; Elevitch, C.R.; Wilkinson, K.M. 2000. Multipurpose trees for agroforestry in the Pacific Islands. Agroforestry 

guides for Pacific Islands #2. Holualoa, HI: Permanent Agriculture Resources. http://www.agroforestry.net/afg/book.html. 

(December 2011).

U.S. Census Bureau. 2014a. International Programs, International 

Database. Midyear population and density: Puerto Rico, USVI, 

American Samoa, Commonwealth of the Northern Mariana 

Islands, Republic of the Marshall Islands, Republic of Palau, 

Federated States of Micronesia, Guam. http://www.census.gov/

population/international/data/idb/region.php (March 2014)

U.S. Census Bureau. 2014b. State & County QuickFacts: Hawai’i. 

Population 2010. http://quickfacts.census.gov/qfd/states/15000.

html (March 2014)

U.S. Congress, Office of Technology Assessment [US OTA]. 1987. 

Integrated renewable resource management for U.S. insular areas. 

OTA-F-325. Washington, DC: U.S. Government Printing Office. 

http://www.princeton.edu/~ota/disk2/1987/8712/871201.pdf. 

(November 2011).

U.S. Department of Agriculture, Forest Service, State and Private 

Forestry [USDA FS SPF], Program Redesign Committee. 2007. 

Tropical forests of the United States: applying U.S. Department 

of Agriculture, State and Private Forestry programs. http://

www.hawaiistateassessment.info/library/tropicalforests-of-theUnited-States-Final60607.pdf. (November 2011).

U.S. Department of the Interior, Bureau of Land Management, 

Seeds of Success [USDI BLM SOS]. 2011. Seeds of success training, module 1: introduction and overview, seed collection for restoration and conservation. Native Plant Materials Development 

Program. Internal document. Washington, DC: U.S. Department 

of the Interior, Bureau of Land Management. http://www.nps.

gov/plants/sos/. (January 2014).

Vieira, D.L.M.; Holl, K.D.; Peneireiro, F.M. 2009. Agro-successional restoration as a strategy to facilitate tropical forest recovery. Restoration Ecology. 17: 451–459.

Withrow-Robinson, B.; Johnson, R. 2006. Selecting native plant 

materials for restoration projects: insuring local adaptation and 

maintaining genetic diversity. Corvallis, OR: Oregon State University Extension Service. http://ir.library.oregonstate.edu/xmlui/

bitstream/handle/1957/20385/em8885-e.pdf. (August 2006).

Wittman, H.; Desmarais, A.; Weibe, N., eds. 2010. Food sovereignty: reconnecting food, nature and community. Oakland, CA: 

Food First Books. 232 p.

Why Start a Tropical Nursery for Native and Traditional Plants? 29
30
Every nursery is unique. Tropical nurseries operate in a vast range of environmental, social, and economic contexts. Each nursery has a unique design based on 

distinct needs, goals, and resources (figure 2.1). With so many diverse factors to 

consider, no standard blueprint for how to design a particular nursery exists. The 

best nursery design will be matched to your specific location, plants, community, 

and goals.

Although planning a nursery can seem like a somewhat complex process, the 

basic formula is to answer the questions of why, who, what, how, where, and when. 

Why start a nursery? Who will the nursery serve? What will the nursery grow? How 

will the plants be propagated and the nursery be managed? Where will the nursery 

be located? When will be best to carry out production tasks? This chapter is intended 

to guide you through the process of answering these questions for planning a native 

plant nursery.

31

Planning a Tropical Nursery

Kim M. Wilkinson and Thomas D. Landis 2

Facing Page: The best nursery design will be matched to your specific context and goals to meet the needs of your clients. 

Photo from American Samoa by Diane Haase.
The Importance of Planning

The startup phase of successful nurseries involves 

thoughtfulness, research, and careful planning. Too often 

this crucial planning phase is rushed or misdirected by 

preconceived ideas of how a nursery should look (such as, 

“all nurseries should have a big greenhouse”) or what the 

nursery should do (such as, “we ought to grow seedlings of 

this species for reforestation”). Before making these types 

of decisions, it is important to do some strategic planning. 

The initial planning phase is an opportunity to step back 

and clarify the vision and goals of the nursery and to coordinate all components needed to reach these goals (figure 

2.2). For example, the first idea may be to build a large 

structure, such as a greenhouse. Deeper exploration of the 

nursery’s goals and the actual needs of the species to be 

grown, however, may reveal how to create a site-appropriate design of several different, smaller scale environments 

that are ultimately more economical, efficient, and effective 

for producing plants.

Planning is essential because each aspect of the nursery 

affects every other aspect. For example, consider the seemingly simple act of choosing what kind of container to 

use for growing plants. Containers come in many sizes 

and shapes (see Chapter 7, Containers) and will dictate 

the nursery layout and benches needed, what types of 

32 Tropical Nursery Manual

Figure 2.1—Native plant nurseries have many unique characteristics, including growing a wide diversity of species. Photo by Tara Luna.

Figure 2.2—Take time to visit other nurseries, talk with potential 

clients, and clarify your vision and goals before starting a nursery. 

Photo by Ronald Overton.
growing media will be used, how seeds will be sown, how 

plants will be watered, how plants will be delivered, and so 

forth. Container type and size will also affect scheduling and 

production costs. It would be unwise to invest in containers without thinking these other factors through. Browsing 

through this handbook can be helpful before making any 

big decisions or purchases. The example with containers is 

only one illustration of the interconnectedness involved in 

planning a nursery. Both direct and indirect factors must 

be considered in nursery design (figure 2.3).

Defining the Nursery’s Vision and 

Objectives

Most nurseries are founded on a vision how the landscape and community could be 10, 50, or 100 or more years 

from now as a result of efforts today. This vision will be a 

guiding force for the nursery’s work.

Some of the vision process involves locating your “north 

star,” the condition in the future that you may be inspired 

to work and move towards your whole life, but may not live 

to see. Visioning is the chance to dream big and may also 

include very personal aspects, such as how you would like 

your lifestyle, livelihood, and role in your community to be.

Any nursery founded to propagate native or culturally 

important plants must be aware of its vital role in protecting ecosystem health and diversity. Biodiversity is defined at 

three different levels (Wilson 1988, Landis and others 1993):

• Genetic level—the number of alleles or genotypes 

within a species.

• Species level—the number of species within a population and the number of these populations within a 

community or ecosystem.

• Ecosystem level—the number of communities and 

ecosystems in the world.

Nurseries play a key role in helping to preserve biodiversity at all three levels (figure 2.4). Mindfulness of appropriate local genetics and diversity allows for nursery work to 

be of enormous benefit for the environment, the community, and future generations of plants, animals, and people.

Determining Community Needs

Visions are translated into practical objectives through 

interactions with the local community and the local environment. The nursery founder may have a vision, for 

example, of seeing many kinds of people planting a wide 

Planning a Tropical Nursery 33

Figure 2.3—A nursery is a web of interrelated factors. Getting a 

good overview of direct and indirect factors for plant production in 

any situation will help design the best nursery to meet local needs. 

Illustration from Dumroese and others (2008).

Figure 2.4—Nurseries play a key role in helping to preserve biodiversity. For example, every time you collect seeds to propagate, 

you can help protect local species and genetic diversity. Photo by 

Douglass F. Jacobs.
diversity of native and culturally important plants provided by the nursery, while native animals from pollinators 

to birds and mammals also benefit. You can hold a broad 

vision for the future, while working practically within a 

scope that makes sense for the present. For example, an 

approach of—“if we grow it, they will plant it”—may result 

in wasted effort if the community has little desire or knowledge about how to use the nursery’s plants. To help bridge 

the gap between a vision and practical objectives, start by 

asking questions such as—

• What is truly needed and wanted in our community?

• Who are the potential clients of our plant materials at 

this time?

• Who might be potential clients in the future (if we 

engage in outreach and education)?

• What are the needs and priorities of the potential 

clients?

Formal and informal avenues should be used to gather 

as much information as possible. Trade groups, guilds, 

elders, and instructors that work with plant products are 

often tremendous sources of information. Holding a gathering, discussing hopes for the future of the local environment and community, interviewing people, publishing an 

article in the local newspaper and asking for responses, or 

conducting formal market research can be useful in this 

phase. It is also imperative to understand the challenges 

people in the community face on diverse outplanting sites 

(figure 2.5). For example, if you plan to grow species to 

serve local farmers for their conservation practices (such 

as erosion control, windbreaks, or riparian buffers), make 

field visits to observe actual conditions on the farms.

This “end-user needs assessment” is key to helping a 

nursery determine not only what species to grow but also 

the container size (stocktype) clients might prefer. With 

culturally important plants, particularly those used for 

medicine and crafts, clients often have exacting specifications. It is essential to assess these expectations during the 

planning phase to ensure that the nursery will provide what 

clients actually need and want (figure 2.6). In addition, 

an assessment will help determine how long the existing 

demand is likely to last, which may indicate whether the 

nursery will be viable in the long term.

The nursery may also use its own vision to avoid being 

swept up into meeting the short-term demands of a changing market. For example, if public interest in planting a 

certain exotic ornamental species becomes high, a nursery 

might expect financial gain from meeting this demand. 

Would meeting that demand, however, help fulfill the 

vision of the nursery? Perhaps it would and perhaps not; 

deciding whether to meet a certain demand depends on the 

circumstances and the nursery’s objectives.

34 Tropical Nursery Manual

Figure 2.5—Understanding the challenges of your client’s outplanting sites is important so you can grow the best plants for their needs. 

A farmer planting trees for forage on a dry site in East Timor (A); “badlands”—bauxite mining areas—on the main island of Palau where 

revegetation is very challenging (B); an abandoned pasture in St. Croix, U.S. Virgin Islands, infested with invasive coral vines (Antigonon 

leptopus) that smother other vegetation (C). Photo A by J.B. Friday, photo B by George Hernández, and photo C by Brian F. Daley. 

B

C

A
Keep in mind that nursery-community communication goes both ways. Although the nursery must listen to 

the needs and wants of its community, it can also share its 

visions and goals by engaging in education and outreach to 

share information with clients and the community about 

the benefits and attributes of its plant materials. For example, if local farmers or landscapers believe only a certain 

ornamental plant works well as a boundary hedge, it may 

be because people are unaware of a native or traditional 

species that can be planted successfully for the same purpose, while providing other benefits. A good understanding 

of local ecology, environmental issues, history, soil types, and 

site needs for outplanting materials is important for effectively communicating with potential clients (figure 2.7).

Defining Target Plant Needs and 

Other Services

Successful nurseries provide healthy, high-quality, 

locally adapted plants that have high survival rates after 

outplanting. In addition, nurseries must provide the fieldready plants for outplanting when environmental conditions and the outplanting sites are optimal for survival. 

Nursery planning is critical for matching the plant materials produced with the needs and conditions of clients’ 

outplanting sites to ensure that the plants can produce 

the materials or products (medicine, wood, food, and so 

on) that clients expect. (See Chapter 3, Defining the Target Plant, for more discussion about this topic.) These 

needs dictate the plant’s target size, age, genetic source, 

container type, and management in the nursery.

Planning a Tropical Nursery 35

Figure 2.6—The perpetuation of culturally important plant materials is a key objective for many tropical nurseries. The nursery must 

work with end-users in the community to ensure that the plants produced will meet cultural needs, such as Kava, an important medicinal 

and ceremonial plant of the Pacific Islands (A) or plants for traditional weaving on Yap (B). Photo A by Ronald Overton, and photo B 

by Megan Parker. 

Figure 2.7—Gathering information about the community 

needs helps clarify what the nursery could grow. Here, villagers in Baucau, Timor Leste, developed a list of local agroforestry 

species and their uses. Villagers used colored markers to vote 

on which trees they would like to plant. The most popular was 

candlenut, Aleurites moluccana, whose nuts could be sold locally 

and processed for food or oil. Ai kameli is sandalwood (Santalum 

album) and ai bubur is Eucalyptus alba, the local firewood. Note 

that the typical forestry trees, teak (Tectona grandis) and toon 

(Toona species), did not receive any votes. The language is Tetun. Photo by J.B. Friday.

A B
Some topics to consider when deciding what to grow 

include—

• The species the nursery might be capable of growing.

• Specific end-user requirements for each species (such 

as medicine, food, timber, habitat, soil stabilization, 

windbreak, and commercial products).

• The types of environments in which plants will be 

outplanted.

• The size and age of stock preferred.

• The season during which people prefer to plant (in the 

tropics, usually the beginning of the rainy season).

• The quantity of each species people may want to 

plant.

• The distance people are willing to travel to obtain the 

plant materials.

Another consideration is what species are most needed 

from an ecological standpoint. For example, in some tropical forests, rarer or later successional species needed to 

restore the diversity of the forest may not return without 

human help, and these species may be a higher priority for 

nursery work (Holl and Aide 2011).

The target plant requirements will differ among species 

and outplanting sites and will influence all aspects of nursery design: location, structures, container types, scheduling, management practices, propagule collection, and so 

forth. In addition to producing plants, many nurseries offer 

a range of services such as consultations and planting plans, 

seed selection and collection, testing of new propagation 

methods, plant delivery, and outplanting support (figure 

2.8). Successful nurseries also dedicate time and resources 

to community education, client communications, marketing, and establishing good relationships with farmers, natural resource managers, landscapers, and planners.

Assessing Resources and Costs

How will we start a nursery? How will the finances 

work? What resources and costs are involved? Is starting a 

nursery even feasible? Each of these questions is discussed 

in the following sections.

Starting With a Small Pilot Nursery

It is often wise to start with a small pilot nursery to better understand how to produce native plants and successfully manage a nursery in your conditions to meet your 

community’s needs. A pilot phase is an opportunity to try 

out production on a smaller scale with less risk (figure 2.9). 

The design of the pilot nursery is essentially your “best 

educated guess” on what type of set-up would be optimal 

to produce plants, based on this handbook and your personal experiences. You can expect a large learning curve 

when developing methods for growing native and culturally important plants. High crop losses may occur during 

the first few seasons, and more losses may take place for 

the sake of experimentation (figure 2.10). A few seasons 

of growth and observation in a pilot nursery can preclude 

unnecessary expenditures, develop viable propagation 

strategies, and eliminate many of the unknowns regarding 

how to produce plants and manage the nursery, thereby 

providing enough detailed information to effectively plan 

a larger facility. A pilot nursery can also be invaluable in 

estimating costs and making a more accurate feasibility 

assessment when the time comes to expand.

36 Tropical Nursery Manual

Figure 2.8—Nurseries dedicate time and resources to cultivating 

relationships with clients, including visiting clients’ sites months 

or years after the plants left the nursery, to see how the plants are 

doing. Photo by Douglass F. Jacobs.

Figure 2.9—Starting with a smaller scale pilot phase reduces the 

number of unknown factors and risks and is a key part of developing a successful nursery. Photo by Thomas D. Landis.
A small pilot nursery can help assess the following 

practices:

• How to take crops through all phases of development, 

from germination through hardening and distribution.

• How to develop realistic timelines and budgets for 

future production.

• How to plan infrastructure for production (and what 

infrastructure is truly necessary versus optional).

• Which growing media, container types, and propagation strategies are optimal for the plants.

• What labor and material costs are involved.

• What types of challenges may arise.

• How clients respond to the plants that were produced.

• How plants perform on different outplanting sites.

• What aspects of the nursery vision are feasible to carry 

out at this time.

The key is to start small, learn lessons along the way, 

and adjust your system accordingly. Keep in mind that 

no one who works with plants will ever feel as if they have 

learned everything they need to know; even very established nurseries are always learning more about how to 

grow and perpetuate plants, how to manage facilities and 

resources better, and how to improve relations with clients 

and the community. At some point, however, you will be 

confident that it is time to expand on the successes of the 

pilot nursery and continue development on a larger scale.

Assess Finances

Finances are an essential part of determining how a 

nursery will start and operate. Nurseries differ greatly in 

terms of their financial objectives. Some nurseries may 

be funded through grants or government programs while 

others are aligned with a particular project or organization. Some may have startup or pilot phase money, but are 

expected to be financially self-sufficient in the future. Private, for-profit nurseries must earn enough income from 

the sale of plants to at least pay for development, infrastructure, production costs, and staff time. Whatever the 

circumstances, finance is a key part of nursery planning. It 

determines—

• How much money can be invested in the nursery at 

the outset.

• If staff can be hired.

• In what timeframe the nursery can start to produce 

plants for sale.

• How many plants can be produced.

• What price can be charged for the plant materials.

Even if plants are to be distributed freely and not sold, 

it is still essential to know the cost to produce each kind of 

plant for planning, assessing feasibility, and ensuring the 

financial viability of the nursery. For a new nursery, predicting the cost of plants is complicated and depends on 

infrastructure, nursery size, staff skills, knowledge base, 

and many other factors. Nurseries that have gone through 

a pilot phase probably have a good grasp of the production costs. These costs can be revised to reflect production 

on a larger scale. Without a pilot phase, estimating costs 

before production is very difficult to do accurately, especially because high crop losses may be expected during the 

first few seasons while successful propagation methods are 

developed.

Visiting other nurseries to get an idea of similar production processes can be very helpful as resources and 

finances are assessed. Government nurseries are a great 

source of information because staff members usually share 

production details openly, and their production costs are 

public information. Visiting private nurseries is also useful in assessing all the stages that go into plant production, 

although they may be less open regarding financial and 

operational details. These production details can then be 

used to estimate costs in the planned nursery.

Estimate crop production costs by considering all phases 

of production, from collecting seeds to mixing growing 

media to delivering the plants (figure 2.11). To improve 

the accuracy of the estimated costs, consider the timeframe 

Planning a Tropical Nursery 37

Figure 2.10—The pilot phase is the time to test ideas and accept 

the fact that failures lead to better results. These growers tried to 

repurpose Styrofoam® cups as nursery containers. They found that, 

unlike proper root-training nursery containers, the round shape 

allowed roots to spiral, which could be a serious problem after outplanting. Photo by J.B. Friday.
for growing the crop, the size of the stock, the labor and 

materials required, and the fact that some crop losses will 

take place during production. Remember to account for the 

following factors:

• Material production costs (for example growing 

media, water, fertilizer, seeds, pest control).

• Labor costs for production, maintenance, and delivery.

• Shipping costs and shipping time required for supplies and equipment (shipping can be costly and slow 

in remote areas and islands—order well in advance).

• Time and labor for customer relations (such as helping clients determine their target plant needs, answering e-mail messages or telephone calls, and visiting 

sites after outplanting).

• Inventory required (for example, the time, space, 

and materials each crop will require, such as nursery 

benches, containers, and trays).

• Structures (for example, protected germination area, 

shadehouse, and storage).

• Overhead costs (such as rent, insurance, water, utilities, and taxes).

• Time and labor for administration (bookkeeping, 

payroll, and staff relations).

• Time and funds for outreach, advertising, or educational programs.

It is wise to estimate a range of best-case (most economical) and worst-case production scenarios. After production 

is underway and actual costs are known, it is imperative to 

revisit the price structure of the plants to ensure that they are 

in line with actual costs. In some cases, the costs of producing 

plants on a larger scale will be lower per plant than during 

the smaller scale pilot phase. But it is also possible that costs 

related to rent, utilities, labor, and so on may be higher.

Assess Feasibility

After assessing resources and costs, it is time to evaluate 

whether starting the nursery is a realistic and achievable 

undertaking. Again, starting with a small pilot phase is a 

good way to determine the feasibility of starting a nursery. The feasibility assessment needs to include the species potentially available to grow, client needs, production 

costs, market price, and the nursery’s site, goals, and capabilities. Will the emphasis be on growing plants from seeds 

or cuttings? How long does it take to grow these species 

to target specifications? What size plants need to be produced? Several scenarios need to be examined, including 

a variety of facility designs, sizes, and locations, so that the 

best conditions to meet projected needs are identified.

Other feasibility questions that need to be asked include, 

“Can the vision and objectives be fulfilled without starting a 

new nursery? Do alternatives exist?” Acting as a distributor 

instead of as a producer may be an economical alternative 

to starting a new nursery; this alternative has its benefits 

and drawbacks (table 2.1).

Selecting a Nursery Site

After the decision has been made to move beyond the 

pilot phase and develop a nursery, an appropriate nursery 

site must be selected. If possible, the pilot nursery can be 

located on the proposed permanent site to maximize familiarity with production and management on that site. Sometimes the nursery site is selected after the pilot phase, and 

this approach also has merit because you may be clearer 

about what you are looking for in a nursery site. Nursery 

site selection involves working with nature, rather than 

against it, for the most effective, efficient, and economical design (figure 2.12). The less the natural environment 

has to be modified to produce high-quality plants, the less 

expense the nursery will have to incur to create optimal 

crop conditions. Again, an understanding of the methods 

and costs associated with producing target plants along 

with the client-needs assessment will help in choosing a 

site. Careful observation of site conditions and an assessment of past and present climatic records are important. 

Even in cases where the site is already chosen, the process 

of site observation and inventory described here is helpful 

to understand the strengths and challenges of that site. Of 

course, no site is perfect.

38 Tropical Nursery Manual

Figure 2.11—To accurately estimate the time and expenses 

required for growing plant materials in a nursery, examine all 

aspects of plant production, from collecting seeds to delivering 

plants. Photo by Thomas D. Landis.
Planning a Tropical Nursery 39

Benefits Drawbacks

Purchase plants

• Time and capital available for other uses

• No nursery staff needed

• More long-term flexibility

• Short-term or no commitment required

• No control over growing process

• Less control over plant quality, genetics, and availability

• Plants may not be adapted to local environment

• Unique needs of local clients may not be met

Start own nursery

• High control over growing practices (can choose organic methods, 

local resources, and so forth)

• High control over quality, genetics, and availability of plants

• Plants will be adapted to local environment

• Can develop local expertise on plant growing and handling

• Can use traditional or culturally appropriate methods if applicable

• No reliance on others to provide plants

• Create job opportunities

• Large initial investment, capital, and time

• Long-term professional and economic commitment

• Native plant markets are notorious for year-to-year fluctuations

• Steep learning curve to gain proficiency in quality plant production 

and nursery management

Table 2.1—The benefits and drawbacks of either starting a nursery or distributing plants from another supplier. Adapted from 

Landis and others (1994).

Figure 2.12—Good site assessment and an understanding of crop needs will help determine the best structures for a nursery. 

Shown here is nursery construction in American Samoa. Photo by Thomas D. Landis.
Critical nursery site selection factors include the following—

• Access to good-quality, affordable, and abundant 

water.

• Unobstructed solar access.

• Easily accessible flat area for delivery or processing of 

bulky, heavy material such as soil, sand, mulch, and 

fertilizer.

• Easy access and close proximity to staff.

• Adequate land area.

• Reliable energy supply (if water pumps, fans, or lights 

are used).

• Freedom from insurmountable ecological concerns 

(such as neighboring chemical pollution, unmanageable noxious weeds, and so on).

• Freedom from problematic political concerns (such as 

problematic zoning and historical land use issues).

Climatic and biological attributes top the list for importance in site selection. An abundance of good-quality, reliable, affordable water is the number one factor; water quality 

always needs to be tested when a site is being considered for 

nursery construction. See Chapter 11, Water Quality and Irrigation, for more information about this topic. Unobstructed 

solar access is also essential; plants need sun to grow. Access 

to electricity is important only if the nursery uses practices 

such as electric water pumps, fans to move air through the 

nursery, lights at night, or communication devices. At least 

one person needs to have quick access to the nursery in case 

of emergency; if the nursery site will be far away from human 

dwellings, it may be advisable to construct a caretaker residence on site. The amount of land selected for the nursery 

must be large enough for the production areas and any support buildings, and also allow for the efficient movement 

of any equipment and materials. In addition to immediate 

needs, potential nursery sites need to be evaluated on the 

basis of available space for possible expansion. Ecopolitical 

site selection factors, notably land use zoning and concerns 

about neighboring land uses that may involve herbicide, 

pesticide, or potential groundwater contamination are also 

important factors for determining suitable sites for nursery 

development.

40 Tropical Nursery Manual

Site 

selection 

criteria

Weight 

valuea

Site A Site B Site C

Rating Weighted 

score Rating Weighted 

score Rating Weighted 

score

Critical Factors

Good solar access 10 9 90 7 70 9 90

Water quality 9 9 81 7 63 4 36

Water quantity 8 10 80 8 64 9 72

Available energy 8 9 72 9 72 10 80

Adequate land area 7 8 56 8 56 10 70

Zoning restrictions 7 10 70 6 42 8 56

Pollution concerns 6 9 54 7 42 9 54

Secondary factors

Microclimate 6 9 54 8 48 9 54

Topography 5 10 50 9 45 10 50

Labor supply 4 9 36 8 32 10 40

Accessibility 4 8 32 6 24 8 32

Shipping distances 3 9 27 7 21 10 30

Total score 702 579 664

Site suitability #1 #3 #2

Table 2.2—Decision matrix for evaluating potential container nursery sites. In this example, Site A received the highest score and is 

therefore considered the best choice for a nursery site. Adapted from Landis and others (1994).
Protected microclimate can make dramatic improvements in nursery productivity and reduce cost expenditures. A site with a climate sheltered from extremes 

such as high winds, storms, and severe temperature fluctuations is ideal. Gentle topography also makes nursery 

set-up and management much easier than hilly or steep 

terrain. Access to the nursery by staff and clients is also 

useful for economical nursery production. In areas at 

risk of high winds, making use of a natural windbreak 

(figure 2.13) or having the ability to quickly remove the 

plastic from the roof of a greenhouse may save a structure in a severe storm. A backup water supply ensures 

crop survival through periods of drought or uncertainty. 

Firebreaks or a site selected to minimize fire risks can 

preclude disaster.

Each potential nursery site will likely have good and 

bad attributes. A decision matrix (table 2.2) can be constructed by listing the potential nursery sites across the 

top and the significant site selection criteria down the 

side. The next step is to assign each site selection criterion 

an importance value on a scale from 1 to 10, with the 

most critical factors receiving the highest values and the 

less important ones receiving progressively lower values. 

Next, the suitability of each potential nursery location is 

evaluated and rated, again on a scale of 1 to 10, based 

on the information that has been gathered. The score for 

each cell in the matrix is then calculated by multiplying 

the weights for each site selection factor by the rating for 

each site. Finally, the weighted scores are totaled for each 

site, and, if the weights and rankings have been objectively assigned, then the potential nursery site with the 

highest total ranking should be the best choice. If all the 

potential sites are close in score, then the process needs 

to be repeated and careful attention paid to the relative 

values of each factor. The selection criteria can also be 

adapted or expanded to include other factors important 

to you. If the scores are still close, the sites are probably 

equally good.

Planning a Tropical Nursery 41

Figure 2.13—Planning for local environmental conditions and for risk is essential during the process of nursery design. This nursery in 

Guam uses natural windbreaks to reduce water use and add protection to the nursery site (A); this nursery in Palau catches rainwater as 

a backup water supply (B). Photo A by Tara Luna, and photo B by George Hernández.

A B
Planning the Flow of Work

Good nursery planning takes place in time and in 

space (figure 2.14). Overall nursery planning examines 

the flow of work and materials through all seasons and 

phases of growth.

The requirements to produce target plants in time for 

the client’s outplanting season guide all other aspects of 

nursery design. Good site selection and a sound knowledge of propagation methods are important for creating 

the most appropriate environments for the crop’s needs. 

Rather than a single, large nursery structure or area, it 

is common to have a diversity of smaller areas and some 

structures tailored to meet the needs of the crops as they 

go through their development.

Although crop production is the core of nursery activities, it is only part of the whole picture. Preparation, 

cleanup, and storage must also be well planned. Where 

will seeds be cleaned, stored, treated, and tested? Where 

will containers be cleaned, sterilized, and stored? Where 

will clients be met? The flow of work through time and 

seasons and design to facilitate the movement of people 

and plants in an efficient and safe way is an essential 

component of nursery planning (figure 2.14).

Considering the nursery’s local effect is also important. 

For example, consider not only where good-quality water 

will come from for irrigation, but also where the water 

could go after nursery use. The water may contain fertilizers and be a potential source of pollution, possibly 

creating problems and legal issues for the nursery. With 

good planning for flow over time, however, that same 

water may be used as a resource, directed to other crops 

(figure 2.15), or recycled. Thoughtful design minimizes 

the amount of water used, provides for the needs of 

plants, and deals with runoff appropriately, as discussed 

in Chapter 11, Water Quality and Irrigation. Consider as 

many elements as you can during the planning phase, and 

be prepared to adapt as new opportunities for efficiency 

present themselves after the nursery is operational. 

Planning as an Ongoing Process

The initial planning phase is a crucial part of successful nursery development, but the planning process does not 

stop after the nursery is operational. The vision of the nursery needs to be revisited regularly. Set aside time to assess 

the nursery’s progress in fulfilling its objectives, visualize 

new possibilities, and adapt to changing circumstances. 

42 Tropical Nursery Manual

Figure 2.15—Good planning translates to making efficient use 

of resources and space. This small greenhouse saves water and 

labor by growing sedges and wetland grasses on benches beneath 

those for dryland forbs. Photo by Tara Luna.

Figure 2.14—Good planning examines the flow of work and sets 

up effective workstations for efficient and enjoyable plant production during all phases of growth. Photo by Kim M. Wilkinson.
Continually learning more through nursery visits, conferences, and field days will also help with ongoing planning 

and improvement (figure 2.16). Following up with clients 

and revisiting target plant specifications are also essential 

for planning for success. See Chapter 18, Working With 

People, and Chapter 19, Nursery Management, for more 

discussion about these topics.

References

Holl, K.D.; Aide, T.M. 2010. When and where to actively restore 

ecosystems?  Forest Ecology and Management. 261(10): 1558–

1563.

Planning a Tropical Nursery 43

Figure 2.16—Planning a nursery is an ongoing process that is aided by visiting other nurseries and continually learning more. Photo 

by Kim M. Wilkinson.

Landis, T.D.; Lippitt, L.A.; Evans, J.M. 1993. Biodiversity and 

ecosystem management: the role of forest and conservation nurseries. In: Landis, T.D., tech. coord. Proceedings, Western Forest 

Nursery Association, 14–18 September 1992—, Fallen Leaf Lake, 

CA. Gen. Tech. Rep. RM-221. Fort Collins, CO: U.S. Department 

of Agriculture, Forest Service, Rocky Mountain Forest and Range 

Experiment Station: 1–17.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1994. 

The container tree manual: volume 1, nursery planning, development, and management. Agriculture Handbook 674. Washington, DC: U.S. Department of Agriculture, Forest Service. 188 p.

Wilson, E.O., ed. 1988. Biodiversity. Washington, DC: National 

Academies Press. 521 p.
44
45

People planting native and traditional species in the tropics have diverse goals and 

face a multitude of challenges on their outplanting sites. Understanding these goals 

and challenges and the steps necessary to meet them is essential to outplanting 

success. The Target Plant Concept is used to define what plant materials to grow in 

the nursery to meet the needs on the outplanting sites and achieve project objectives. 

A “one-size-fits-all” approach to plant materials does not work well. Instead, plant 

materials must be matched to meet the challenges of the outplanting site. Establishing 

targets is critical; for a given species, the target plant destined for a harsh restoration 

site where care will be minimal after outplanting is very different from one that will 

be outplanted in a city park or hotel landscape where it will be pampered. Thus, the 

definition of the target plant depends on how it will be used—its “fitness for purpose” 

(Sutton 1980). This definition is the essence of the Target Plant Concept.

The Target Plant Concept was developed as a way to define the target plant, and 

it has two equally important components. The first component incorporates three 

simple approaches that provide the broad, fundamental basis necessary to successfully 

complete outplanting projects (figure 3.1). The second component is the step-by-step 

process of defining the target plant material. The nursery manager and client systematically answer eight sequential, but interrelated questions about the outplanting site 

and project goals to ultimately define the target plant material. The target plants are 

then produced, outplanted, and monitored; then the cycle repeats. An understanding 

of the Target Plant Concept is vital when starting a new nursery or upgrading an existing one and is also useful when working with customers. Let us examine the first of the 

two components in detail.

Defining the Target Plant

Thomas D. Landis and Kim M. Wilkinson 3

Facing Page: The “target plant” is cultivated to meet the challenges of the outplanting site and the objectives of the 

project. Photo by J.B. Friday. 
Important Approaches for 

Implementing the Target 

Plant Concept

The first component of the Target Plant Concept incorporates three simple, often overlooked ideas that, when 

considered together, guide the broad approach for defining 

and selecting the target plant materials for a specific site.

Start at the Outplanting Site

With the Target Plant Concept, the nursery production process starts with the characteristics of the outplanting site. Land managers specify exactly what type 

of plant material would be best for their site conditions 

(figure 3.2). Understanding these needs, the nursery 

grows plant materials that are locally adapted, genetically appropriate, and the optimal size, age, and so on to 

survive and thrive on the outplanting site.

Forge a Nursery-Client Partnership

With the Target Plant Concept, the land manager and 

nursery manager work together to define the ideal type 

of plant for the project, the nursery grows the plants, and 

they are outplanted (figure 3.3). Based on performance 

after outplanting, the land manager and nursery manager may work together to revise target plant characteristics to improve survival and growth of future crops 

(figure 3.4). See Chapter 18, Working With People, for 

information on how to facilitate meaningful, mutually 

supportive dialogue after outplanting. Good communication between native plant customers and nursery 

managers builds partnerships and ensures the best possible plants for the project. This feedback also ensures 

that both nurseries and clients are learning and adapting, 

thereby improving successes over time.

46 Tropical Nursery Manual

Figure 3.1—The Target Plant Concept starts with a partnership between the client and nursery manager that focuses on putting the 

best plant materials on specific project sites. The manager and client then answer eight questions about the project; their answers define 

the target plant material necessary to meet the objectives of that project. The nursery produces the plants. The client and manager 

subsequently reassess successes and failures and use that information to improve the next crop. Adapted from Landis (2011).
Defining the Target Plant 47

Figure 3.2—Plant materials must be matched to meet the challenges of the outplanting sites and the goals of the projects they serve. 

A nursery in East Timor grows agroforestry species in polyethylene bags to serve village needs (A). The Palau Municipal Nursery grows 

native and culturally important seedlings in large pots for urban forestry projects (B). Future Forests Nursery in Hawai‘i grows native 

trees and shrubs in several types of containers and growing environments for diverse restoration and farm forestry projects (C). Photo A 

by J.B. Friday, photo B by Katie Friday, and photo C by Jill Wagner.

Figure 3.3—Using the Target Plant Concept, nursery and land managers work together to specify what type of plant material would 

be best suited for the project. Shown is a native reforestation project in Guam, where target plants included native trees such as Intsia 

bijuga grown in large, 1-gallon root-training square containers (A). The native seedlings were planted using a mattock in soft clay soil 

(B). Photos by J.B. Friday.

Figure 3.4—Using the Target Plant Concept, the land 

owner and nursery manager work together to define the 

ideal type of plant for the project. Based on performance 

of the first crop, the managers make necessary changes to 

improve survival and growth. Here a nursery manager 

and land owner are standing in a 1-year-old mixed forest planting (foreground) and discussing performance 

of a 4-year-old planting (background) to make some 

decisions together about next year’s target plants. Photo 

by Craig R. Elevitch.

A B C

A B
Emphasize Seedling Quality,

Not Appearance

With the Target Plant Concept, plant quality is not 

determined by how good a plant looks as it sits in the nursery, but by outplanting performance. A beautiful crop of 

plants in the nursery may perform miserably if the plants 

are inappropriate for conditions on the outplanting site. 

Without the Target Plant Concept, inexperienced clients 

may believe they can find cheap, all-purpose native plants 

that will thrive nearly anywhere. Using the Target Plant 

Concept, plants are grown in the nursery with the view of 

their fitness to thrive on the outplanting site and their ability to fulfill the project objectives.

The second component of the Target Plant Concept is the 

process of defining the target plant materials. The nursery 

manager and client use the characteristics of the outplanting site to systematically answer eight sequential, but interrelated questions (figure 3.1) to ultimately define the target 

plant material.

1. What Are the Outplanting

Objectives?

Native and traditional plants are grown for a variety of 

reasons; these project objectives critically influence target 

plant characteristics (figure 3.5).

In the tropics, project objectives may include reforesting 

land that has been deforested; enriching sustainable agriculture with windbreaks or erosion control using native species; 

providing shade in pastures; producing timber or wood for 

crafts; rekindling traditional agroforestry practices; ensuring local supplies of cultural or medicinal plants; restoring 

degraded land; controlling invasive species; creating habitat for pollinators, birds, or wildlife; planting native trees 

in urban areas; protecting and producing traditional foods; 

conserving soil and water quality; educating young people; 

and developing small businesses. Some projects hold a combination of objectives. 

48 Tropical Nursery Manual

Figure 3.5—Native and traditional plants are grown for a variety of project objectives, which 

critically influence target plant 

characteristics. Objectives may be 

production of teak for commercial timber in Samoa (A); regeneration of heritage species, such as 

breadfruit trees, for food security 

(B); perpetuation of native and 

traditional species to sustain traditional lifeways and craftspeople 

(C); and reforestation of sites 

overcome by invasive species, such 

as this swordgrass area on Guam 

(D). Photos by Thomas D. Landis.

B

A C

D
See the following examples of how target plants are linked 

to project objectives:

• Commercial timber production is a common objective 

for planting trees. For this objective of producing saw 

logs, the target plant would be a commercially valuable 

tree species that has been genetically selected for fast 

growth, good form, or desirable wood quality.

• The target plant for a watershed restoration project 

to stop erosion, stabilize the stream bank, and ultimately restore a functional plant community could 

be riparian trees and shrubs with extensive root 

systems and thick stems to withstand flowing water.

• Projects restoring threatened or endangered native 

plant species require target plants that perpetuate the genetic diversity and unique adaptations of 

these imperiled local populations or may require 

target plants that can restore or create critical habitat for these ecologically important species.

• Projects to restore or enhance food security and 

food sovereignty might require target plants that 

are native foods and cultivars of traditional or heritage food plants and varieties. Target plants may 

also include species and genotypes that yield the 

most or best-quality fruits, nuts, or other food.

• Project objectives for a burned, eroding former 

sugarcane field might be to stop soil erosion, 

replace exotic weed species with native or other 

desirable plants, restore nutrient cycling, and build 

up organic matter for future productivity. Target 

plants for such a project might include a direct 

seeding of native grasses and forbs or possibly a 

noninvasive annual plant to first stabilize the soil, 

followed by an outplanting of native nitrogenfixing trees and other species to out-compete the 

weeds and build fertility on the site. These actions 

can improve site conditions over time. Later, more 

diverse species can be planted.

• Renewing native and traditional species for crafts or 

other cultural uses is an objective for many projects. 

Target plants may include local cultural plant species to 

help protect wild plants from the stresses of overcollection and to make their collection and use 

more accessible to elders and young people.

• Some projects have the objective to reduce the effects 

of weeds or invasive species. The presence of invasive 

exotic plants is often a result of disturbances that create 

ideal conditions for these plants to thrive. Therefore, 

simply removing undesired species does not solve the 

problem. Instead, undesirable plants must be replaced 

with desired ones. For example, most invasive grasses 

thrive in full sun, so the target plant to shade out the 

grass would need to be large enough to overtop the 

grass immediately after outplanting, and quickly form 

dense shade.

Sometimes project objectives are straightforward and 

target plants are easily defined. For complex, large, or specialized projects, such as Forest Stewardship, conservation, 

or habitat improvement, the assistance of a professional 

may be needed. This expertise helps define objectives and 

target plant material requirements for every species to be 

planted. Many restoration projects follow an approach that 

includes finding reference sites, considering succession, 

and creating measurable goals.

Selecting Reference Sites

“Reference sites” are natural or recovered areas that 

serve as models for desirable recovery of native plant 

communities (figure 3.6A). The comparison of the soils, 

climate, vegetation, and other characteristics of reference 

sites to the project site provides guidance about what species may be established and is essential for setting attainable, site-appropriate goals for restoration (Steinfeld and 

others 2007) (figure 3.6B). Often several reference sites of 

different ages and recovery stages are used for one project. Reference sites also show succession, which is how 

plant communities may change over time.

Planning for Succession

Land managers need to understand not only how a healthy 

200-year-old forest looks, but also how a healthy recovering 

tropical forest looks 1, 3, or 5 years into its development. 

Understanding this recovery process is necessary because 

if native species can colonize and become established on a 

disturbance, the processes of succession (ecosystem development over time), including soil genesis and nutrient cycling, 

are initiated (figure 3.7). Effective revegetation of disturbed 

sites aims to initiate or accelerate processes of natural succession following disturbances. Native plants may be established on disturbed sites through seeding or planting (figure 

3.8). Sometimes passive revegetation (natural colonization) 

of some species is possible where native seed banks are available and limiting factors are mitigated. If passive restoration 

is possible, nurseries may be involved in growing enrichment or later successional species that will not regenerate on 

the site naturally.

Translating Objectives into 

Measurable Goals

Some projects formally translate their objectives into 

measurable goals within specified time frames. These goals 

Defining the Target Plant 49
are sometimes called success criteria or desired future conditions. For example, a reforestation project might set a 

goal of 400 living trees per acre 2 years after outplanting 

whereas a native plant project may have a goal of 75-percent vegetative ground cover of which 90 percent will be 

composed of perennial native species 1 year after outplanting. Production of food, craft materials, or timber might 

be stated in yields. Habitat restoration will define desired 

fauna moving into the planting area and will provide information about the plants themselves. Many restoration 

projects take place in stages. Short-term objectives might 

be to stabilize the soil and reinitiate natural succession, 

whereas longer term objectives include greater diversity 

and complexity. Monitoring is then planned to measure 

if these goals were achieved. See Chapter 17, Outplanting, for more information.

50 Tropical Nursery Manual

Figure 3.6—Reference sites provide guidance about setting attainable, site-appropriate goals for restoring the outplanting site, including which species to plant. Information about conditions on reference sites is compared with the conditions on the outplanting 

site. Measuring trees on a regenerating, 30-year-old stand of native forest after cattle were removed from a pasture (A); surveying a 

pasture where forest recovery is desired (B). Photo A by J.B. Friday, and photo B by Douglass F. Jacobs.

Figure 3.7—Effective revegetation of disturbed sites aims to initiate or accelerate processes of natural succession following disturbances. 

Reference sites of different ages (5 years recovered, 20 years recovered, and so on) help to set reasonable timelines and appropriate species 

compositions for project development. Illustration adapted from Cerro Nara Rainforest Conservation (2010) by Jim Marin.

A B
2. What Are the Conditions of the 

Outplanting Site?

Site evaluation is necessary for clients to determine 

their target plant requirements and achieve their goals. 

The information-gathering step may be informal, based 

on observation and spending time on the site. More complex projects often involve different tests and a systematic approach to determine site conditions. The most 

elaborate site assessments are usually carried out to meet 

regulatory requirements, such as a project requiring an 

environmental impact statement before it can proceed. 

For restoration of disturbed sites, the reference sites 

described previously are also surveyed to compare their 

conditions with those of the outplanting sites. At a minimum, basic land features need to be mapped and the site 

evaluated for basic information about soils, vegetation, 

climate, and site history.

Maps

Finding or creating a basic site map is an important step 

for projects of any scale. Topographic maps and aerial photos 

are especially useful and usually easy to find. Topographic 

maps indicate elevations, contour lines, waterways and water 

features, slopes, aspects, and human-made features including roads and settlements. Aerial photos can reveal features 

such as vegetation, access, structures, and surroundings and 

offer a snapshot of the conditions at one point in time. Special maps can be obtained as needed for more detailed information such as hydrology, soils, or forest cover.

Soils

The U.S. Department of Agriculture, Natural Resources 

Conservation Service (NRCS) and other organizations have 

soil surveys available for many areas; these surveys provide 

basic information on soil type and slope. In addition, a soil 

test can provide information on pH and nutrients (the client can contact the extension service to find out how to supply a soil sample to a testing facility). The client’s personal 

observations are also a valuable addition. From these tests 

and observations, challenges such as issues with drainage, 

compaction, erosion, incorrect pH, depth to water table, 

and so on can be identified. For projects on large areas, a 

soil scientist may be called in to survey the site and provide 

a report, usually in combination with the NRCS or other 

data.

Vegetation

Vegetation information is obtained by observing and 

recording the site’s existing vegetation. At its simplest, 

clients walk their site and list the vegetation they see, 

getting some help from a plant-savvy friend for species 

they cannot identify. Often, publications are available to 

Defining the Target Plant 51

Figure 3.8—At Hakalau Forest National Wildlife Refuge, koa (Acacia koa) seedlings are outplanted in three parallel lines marching up 

the hill from the lower forest to the upper pastures (A). The goal is to establish corridors so that the forest birds can increase their range 

uphill. Scientists at the refuge monitor the health and population of the native forest birds, such as this ‘Apapane (Himatione sanguinea) 

(B). Photos by J.B. Friday.

A B
help identify plants and plant communities. More detail is 

reached when botanists survey the entire site and create a 

comprehensive list of species present, categorized as native, 

desired, or problematic, and sometimes further defined by 

successional phase.

When Global Positioning System (GPS) receivers are 

used for surveys, all the plant information collected is spatially explicit and mapping, analyzing, and sharing data 

becomes easier. Rapid Ecological Assessments are increasingly common and include vegetation mapping at various 

spatial scales. The advantage to this tool is that the data has 

a spatial component and the vegetation data is easily combined with soil and topography in a Geographic Information System (GIS).

Observing the vegetation also verifies and reveals additional characteristics of the site: dry areas, bogs, and areas 

subject to strong prevailing winds. Understanding the 

current vegetation is essential for site-appropriate actions 

later, including protecting and perpetuating native species 

already on the site.

Climate

Climate information can be obtained from national (for 

example, the National Oceanic and Atmospheric Administration) or regional weather service and charted back in 

some places for decades. At a minimum, clients need to 

understand the average rainfall and temperatures on their 

site and the fluctuations and patterns of these conditions 

throughout the year. For example, trends of when the rainy 

season normally starts and ends and the intensity of rain 

events are as important, if not more important, as only 

knowing the average annual rainfall.

Climate data is too broad to use on its own without also 

observing the microclimates on the site. Two sites situated 

only a few miles apart can experience dramatically different 

wind effects, rainfall, and temperatures (Mollison and Slay 

1991). Onsite observation can help the client determine key 

climate information including aspect, winds, and moisture.

Aspect is whether a slope faces the sun (facing south 

above the equator or north below the equator) or away from 

the sun. Sun-facing slopes are warmer and drier; slopes facing away from the sun are cooler and wetter. Aspect is a 

major determinant of vegetation types on a site.

In most tropical areas, especially islands, storm winds 

and hurricanes may come from any direction, but prevailing winds are consistent and can be anticipated (Mollison 

and Slay 1991). Observing vegetation, especially trees bent 

or flagging in one direction, also indicates the direction of 

prevailing winds.

Water and the way it moves through a site are important factors for microclimate. Salt spray, fog, and mist can 

influence site conditions. The site’s proximity to the ocean 

will determine the extent of the moderating effects of the 

ocean. Water features such as ponds, lakes, springs, and 

streams also create microclimates that affect vegetation.

Site History

Site history can be as simple as asking the elder neighbors 

about the past uses and conditions of the site, or as elaborate 

as a full archeological survey (required for some projects). An 

intermediate option is going to the local library or to the Internet to find aerial and other photos of the site dating back over 

time (figure 3.9). Cadastral offices provide publically available 

information on zoning changes, construction, and ownership 

changes of parcels. Information from the soils, vegetation, 

and climate surveys can be used to understand site history 

including floods, landslides, fires, severe storms, and other 

events that took place in the past and may take place again.

3. What Factors on the Project 

Site Could Limit Success?

The information gathered answering question 2 is used 

to identify the environmental factors that are most limiting 

to plant survival and growth on the site and to specify which 

plant species and stocktype would be most appropriate (figure 

3.10 and table 3.1). On most outplanting sites, newly planted 

plants must quickly establish root contact with the surrounding soil to obtain enough water to survive and grow. Water is 

often the most limiting factor, especially on sites with a pronounced dry season or low annual rainfall. Where populations 

of grazing and browsing animals are high, animals may be the 

52 Tropical Nursery Manual

Figure 3.9—Native tree growers and their partners consult aerial photographs and discuss elevation, exposure, and other site conditions in preparation for a forest restoration project in St. Croix, 

U.S. Virgin Islands. Photo by Brian F. Daley.
most limiting factor because they will eat any plants you try 

to establish. If wildfire is a recurring disturbance on the site, 

then soil conditions are often severely altered and very specific measures, such as fire breaks are essential in the early 

years of the restoration project. Severe soil disturbances can 

also eliminate all soil microorganisms including mycorrhizal fungi. Therefore, plants destined for severely altered 

sites need to be inoculated with the appropriate symbionts 

before outplanting. (See Chapter 13, Beneficial Microorganisms, for a complete discussion on this topic.) Riparian 

restoration projects may require bioengineering structures 

to stabilize streambanks and retard soil erosion before the 

site can be planted (Hoag and Landis 2001). In desert areas, 

low soil moisture, hot temperatures, and high winds with 

sand blast may be listed as limiting factors. High winds, salt 

spray, weeds, soil fertility, seasonal flooding, insect pests, 

and land use challenges, such as people driving over restoration areas, are further examples of limiting factors.

Awareness of native plant needs and ecology (based on reference sites) is important to determine if the factors are truly 

limiting based on project objectives. For example, a site may 

not receive much rainfall, but if the goal is dryland ecosystem 

restoration and the dryland native species are adapted to that 

amount of rainfall, these plants will be able to thrive on the site.

4. How Will Limiting Factors 

Be Mitigated?

Mitigating measures are the steps the client will take to 

overcome any limiting factors on the site and achieve project 

Defining the Target Plant 53

Figure 3.10—Limiting factors to a given project site can be displayed as unequal boards of a barrel. Water can only be held to the 

level of the most limiting factor. Illustration by Jim Marin.

Table 3.1—Comparison of different plant establishment methods. Adapted from Landis and others (1992).

Characteristics Transplanting 

wildlings

Outplanting 

nonrooted

cuttings

Direct

seeding

Outplanting

nursery stock

Efficient use of seeds and cuttings N/A* No No Yes

Cost of establishment High Moderate Low Moderate

Ability to establish difficult species Yes No No Yes

Option of using specific genotypes No No Yes Yes

Precise scheduling of plant establishment Yes Yes No Yes

Control of stand composition and density Yes Yes No Yes

Matching stocktypes to site conditions No No No Yes

Depletion of adjacent plant stands Yes Yes No No

*N/A: Not applicable.
Figure 3.11—Target plants are cultivated with the limiting factors of the site in mind. “Badlands” on the main island of Palau are bauxite 

mining areas developed before World War II that are still not revegetating (A). Nursery site for native seedlings for restoration of the mined site 

(B). A native species, Calophyllum inophyllum, outplanted on the badlands (C). Photos by George Hernández.

objectives. Some mitigating measures can be defined by the 

client and included in his or her target plant requirements 

(figure 3.11); others must be carried out by the client on his 

or her site to establish plants successfully.

Nursery efforts to produce target plants will sometimes be the key to overcoming certain limiting factors 

on the site. For example, an absence of beneficial microorganisms on the site can be mitigated by inoculating 

plants with beneficial microorganisms in the nursery, 

as described in Chapter 13, Beneficial Microorganisms. 

Sometimes mitigating measures will be a combination 

of nursery efforts to produce target plant materials and 

client efforts on the site (figure 3.12). For example, sites 

with strong prevailing winds may call for target seedlings 

with sturdy, thick stems to withstand the winds; at the 

same time, the client may install wind barriers around the 

trees to help them get established, or windbreaks may be 

planted to protect the area in advance of other plantings.

54 Tropical Nursery Manual

Figure 3.12—Overcoming limiting factors to restore disturbed 

sites involves a combination of nursery and land manager efforts. 

On a site with tall grass, the land manager would take steps to control the grass competition, and the nursery may also grow trees tall 

enough to overtop the grass quickly. Photo by Douglass F. Jacobs. 

B

A C
For sites with browsing or grazing animals, species 

selection for less palatable species may be important, but 

the land manager must find ways to exclude these animals 

from newly planted areas or at least provide shelter for the 

plants as they establish (figure 3.13). Individual plants can 

be protected with netting or fencing or the entire project 

area may be fenced. In some areas, the presence of wild 

pigs or other animals may make restoration impossible, in 

which case, whole restoration areas may need to be fenced 

and managed to exclude problematic animals.

Limiting factors that cannot be mitigated through reasonable target plants and land manager efforts in the field 

require the client to revise project objectives to be appropriate and achievable on their project site.

5. What Species and Genetic 

Sources Will Meet Project 

Objectives?

Land managers decide what species they will plant 

based on project objectives, reference sites, project site 

conditions, and limiting factors as described previously 

(figure 3.14). The reference sites provide a natural model 

for determining appropriate species and plant community 

composition (what percent of which species, spaced how 

far apart) for the recovering site (figure 3.15).

In some cases, mitigation of limiting factors alone is sufficient to allow for natural regeneration (passive restoration) 

of certain native and desirable plant species on the site. For 

example, on some upland sites in Hawai‘i, simply removing 

grazing and browsing animals and scarifying the soil allows 

long-dormant koa seeds in the seed bank to sprout (figure 

3.16). In these cases, the nursery work may focus on those 

species that do not regenerate naturally. On many disturbed 

sites, however, no amount of mitigation will lead to adequate 

natural regeneration of native or desired species, and in 

these cases, the nursery will be involved in propagating these 

species for reintroduction by planting.

Defining the Target Plant 55

Figure 3.13—Goats eat anything—very few target plants could 

be cultivated to co-exist with goats. If goat browsing is a limiting 

factor, the land manager will need to exclude goats with fencing. 

Photo by Kim M. Wilkinson.

Figure 3.14.—A forester in East Timor develops a tree selection chart for different planting sites (along the left) and different uses 

(across the top). Planting sites include near springs and on farm fields. Uses include food, fallow improvement, firewood, and timber. 

Photo by J.B. Friday. 
Of course, most nurseries have the occasional call from 

a person who asks, “What should I plant?” It is not surprising that some potential clients expect the nursery to tell 

them what to plant. Using the Target Plant Concept, the client and the nursery work together to determine what plant 

species are appropriate to plant for a given site.

Genetic Sources

In addition to proper species selection, three factors 

concerning genetics need to be considered when collecting plant materials: local adaptation, genetic diversity, and 

sexual diversity. See Chapter 8, Collecting, Processing, and 

Storing Seeds, for additional information.

Local Adaptation

Plants are genetically adapted to local environmental conditions and, for that reason, plant materials should 

always be collected within the same area where the plants 

will be outplanted. “Seed zone,” “seed source,” and “seed 

lot” are terms used to identify seed collections. A seed zone 

is a geographic area that has relatively similar climate and 

soil type (figure 3.17). For some native species, seed zones or 

transfer guidelines have been defined by geneticists to help 

land managers choose appropriate seed sources. For most 

tropical native species, such work has yet to be completed, so 

growers are advised to use sources from the same geographic 

area, environmental conditions, and elevation in which the 

nursery stock is to be outplanted.

56 Tropical Nursery Manual

Figure 3.16—In some cases, some species may be able to regenerate naturally if limiting factors are removed. This photo shows 

natural regeneration of Acacia koa on an upland site after grazing 

cattle and horses were excluded. Photo by Douglass F. Jacobs.

Figure 3.17—Seed zones have been developed for some species to 

provide guidelines for collecting seed from the same geographic area 

and elevation zone. Where no seed zones have been defined, do your 

best to collect plant materials from the same geographic area, environmental conditions, and elevation in which the nursery stock is to 

be outplanted. Adapted from St. Clair and Johnson (2004).

Figure 3.15—Reference sites 

and succession processes guide 

which species to plant. One of the 

few endangered Serianthes nelsonii trees surviving on Rota (A). 

Because natural regeneration is 

not enough to sustain populations, nurseries on Rota produce 

Serianthes nelsonii seedlings (B). 

Photos by Thomas D. Landis.

A B
Local adaptation can affect outplanting survival and 

growth in many ways, including growth rate and environmental tolerances. For example, in temperate areas, commercial 

conifers grown from seeds or cuttings collected from higher 

latitudes or elevations will grow slower but tend to be more 

cold tolerant than those collected from lower elevations or 

more southerly latitudes (St. Clair and Johnson 2003). Many 

tropical islands have a pronounced “wet side” and “dry side.” 

Conventional wisdom and research (Ares and others 2000) 

indicates to refrain from moving plant materials from the wet 

side of an island to the dry side, or vice-versa. In addition to 

rainfall tolerances, local adaptation issues may be important 

for other site conditions such as soil types.

In some cases, local adaptation may be essential for 

long-term viability and habitat value of restoration plantings. For example, local pollinators are often adapted to 

the flower sizes and shapes of their locally adapted food 

plants (Kramer 2007). If a site is restored with the same 

species, but not the appropriate local adaptation of that 

species, local pollinators may not be able to pollinate 

them, leading to a project that cannot perpetuate itself.

Genetic and Sexual Diversity

Target plant materials should attempt to represent all 

the genetic and sexual diversity present on the reference 

sites (figure 3.18). To maximize genetic diversity, seeds 

or cuttings need to be collected from as many plants as 

possible. Guinan (1993) provides an excellent discussion 

of all factors involved in preserving biodiversity when 

collecting plant materials and suggests collecting from 

50 to 100 donor plants.

Dioecious species are challenging because they have 

male and female plants. Therefore, all vegetatively propagated dioecious plants will be the same sex as their parent, which can be particularly important on sites where 

a male population is geographically separated from a 

female population. In such cases, plant material collectors may not realize they have collected only one sex, 

leading to a single sex being outplanted on a project site. 

The resulting lack of seed production would compromise project objectives and future natural regeneration. 

Therefore, when collecting cuttings from dioecious species, care must be taken to ensure that male and female 

Defining the Target Plant 57

Figure 3.18—When collecting seed or cuttings from plants, genetic and sexual diversity must be considered. Adapted from Landis and 

others (2003).
Figure 3.20—Plant materials must be sourced from parents that 

possess the desired attributes, especially in the case of medicinal or 

food plants. Piper methysticum (kava) is an important ceremonial 

and medicinal plant for many Pacific Island cultures, and different 

cultivars have different properties and concentrations of active compounds (A). The essential food plants Colocasia esculenta (taro) and 

Musa species (banana) have diverse heritage cultivars with a wide 

range of properties and tastes (B). Photos by Thomas D. Landis. 

plants are equally represented. Dioecious species include 

cycads (Cycadaceae, Stangeriaceae, and Zamiaceae families), 

mulberries (Morus species), pandanus (Pandanus tectorius) (figure 3.19), kukui nut (Aleurites moluccana), and 

the rare Caribbean “yellow prickle” (Zanthoxylum species). Rooted cuttings or other vegetatively propagated 

stocktypes need to be labeled by sex so that males and 

females can be outplanted in a mixed pattern to promote 

seed production.

Additional Considerations About Source

To meet project objectives, other selection factors may 

be considered when defining the appropriate propagule 

sources. In special cases, seeds must be collected from parents that possess desired attributes. For example, if a native 

or traditional species is being grown for timber or craft use, 

seeds need to be collected from trees showing the desired 

form or wood characteristics. Plants grown for traditional 

medicinal purposes must meet the end user’s exacting 

requirements for quality and potency of the source plants 

(figure 3.20). When applicable, these other source considerations are in addition to, not instead of, the three factors 

of local adaptedness, genetic diversity, and sexual diversity. 

For traditional species, the genetic situation needs to be 

considered on a case-by-case basis; sometimes perpetuating heritage varieties is the goal and sometimes improving 

the genetic diversity will be the goal.

6. What Types of Plant Materials 

(Stocktypes) Are Best Suited to 

the Project Site and Objectives?

Common plant materials include traditional nursery 

stocktypes, such as container seedlings, bareroot seedlings, and rooted cuttings, as well as wildings, rootstock, 

nonrooted cuttings, and seeds (table 3.2). Appropriate 

stocktypes for a project are determined by considering 

limiting factors, species, and genetics. The characteristics 

of a particular species will help determine if direct seeding, nursery propagation, or other methods are the more 

appropriate strategy (Steinfeld and others 2007). Many 

options can be considered; some options are summarized 

in table 3.1. Native plant nurseries can provide a wide 

variety of plant materials that will meet the needs of any 

reforestation or restoration project (table 3.2).

58 Tropical Nursery Manual

Figure 3.19—Pandanus tectorius, an example of a dioecious 

plant of Polynesia. Photo by J.B. Friday.

B

A
Seeds

Seeds are often easy to handle, store, and outplant, but 

the effectiveness of seeding directly on the project site varies with species, harshness of the site, project objectives, 

and project timeframe. Directly broadcasting seeds offers 

three principal advantages: (1) seeds can be inexpensive 

compared with other plant materials, (2) spreading seeds 

is relatively easy, and (3) seedlings from broadcast seeds 

develop a natural root system and occur in a more random 

(that is, “natural”) pattern.

Many drawbacks exist as well. Even when the correct 

species and origin are located, seeds are (1) often difficult to 

obtain or are very expensive; (2) not produced in adequate 

numbers every year; (3) sometimes require specialized cleaning and processing; and (4) difficult to store. In addition, predation by birds and rodents, competition from weed species, 

and unpredictable weather often reduce establishment success (Bean and others 2004). Finally, with direct seeding, it 

is difficult to control species composition and plant spacing 

over the project area (Landis and others 1992).

Direct seeding is usually most successful for grasses, 

forbs, and some woody shrubs. Seeding with native grass 

species after wildfires is often used to stabilize soils and 

prevent erosion. Some trees may be established through 

direct seeding, particularly those with large seeds. In California, the direct seeding of acorns to establish native oaks 

has been quite successful (Landis and others 1992). Sowing 

seeds directly in the field may result in poor germination 

and survival. Therefore, direct seeding is recommended only 

for species in which efficient use of seeds is not necessary.

Bareroot Plants

Bareroot plants are started from seeds or cuttings, are 

grown in the ground or in raised beds, and are harvested 

without soil around their roots (figure 3.21). Because they 

require a considerable amount of high-quality soil and 

often take longer to reach shippable size, fewer species 

of native plants are grown for conservation and restoration as bareroot stock than are grown for commercially 

important objectives, such as timber production. Tropical soils are challenging to work with, so bareroot plants 

are usually produced in raised beds with a mix of growing media (for example, sand mixed with compost) on 

a drip irrigation line. One serious drawback is bareroot 

stock needs more postharvest care than container stock. 

Bareroot is suitable only for plants with root systems that 

Defining the Target Plant 59

Table 3.2—Many different types of native plant materials can be provided by nurseries.

Plant

materials Examples Advantages Disadvantages

Seeds Grasses, forbs

• Small and easy to outplant

• Seeds of some native plants can be stored for long periods 

• Plants develop natural root structure

• Maintain genetic diversity

• Some species do not produce 

seeds regularly

• Many tropical seeds do not store well

• Direct seeding more inefficient use 

of seeds than nursery plants

Bareroot 

plants

Coconut, 

mahogany, cedrela

• Less expensive to produce in nursery than container plants

• Easier to transport

• Roots have not been restricted by containers

• Take longer to produce

• Roots dry out easily

• Often lower survival and slower 

growth especially on drier sites

Container 

plants All species

• Well-established root systems means less transplant shock

• Available in a variety of sizes

• Can be planted all year long

• Large stocktypes provide “instant” plants on site

• More expensive to propagate

• More difficult to transport, 

especially larger stocktypes

Root stock Yams, kava, 

breadfruit, bananas

• Easy to store and transport

• Excellent survival after outplanting

• Only works with certain species

Nonrooted 

cuttings

Erithrina, Gliricidia, 

gumbo-limbo, 

Guazuma ulmifolia

• Ideal for live stakes

• Can be efficiently and economically produced in nursery 

stooling beds

• Only works with species that root 

easily

• Best for mesic environments

Layer 

cuttings

Citrus, 

breadfruit

• Do not have to rely on seed crops 

• Ideal for maintaining same genotype

• Requires healthy mother plant

• Only works with species that 

root easily

Rooted 

cuttings Many species

• Do not have to rely on seed crops

• Ideal for maintaining same genotype

• Stooling blocks can be developed for large multiyear projects

• Must be handled carefully during 

transportation and outplanting
can tolerate more disturbance and handling than species 

that need their root systems kept in contact with soil during 

harvest, shipping, and outplanting.

Container Plants

Container plants are the stocktype of choice for many 

tropical nurseries. Container propagation is best when 

small amounts of many different native plants are desired. 

Another advantage is that container stock is more tolerant 

and durable during handling, shipping, and outplanting 

(figure 3.22). Trees and shrubs are typically established 

using container stock rather than by direct seeding for two 

reasons. First, obtaining seeds from most tree and shrub 

species is expensive and time consuming; in many years, 

seeds can be difficult to find. Second, shrub and tree seeds 

germinate and grow into seedlings at a slower rate than 

grass and forb species, giving them a disadvantage on sites 

where grasses and forbs are present. Outplanting shrubs 

and trees as plants gives them a competitive advantage over 

grasses and forbs because the plants have access to full sun 

and roots are often longer and better developed, allowing 

access to deeper soil moisture. In general, grass and forb species are seldom established from container plants because of 

the high cost. Exceptions are when grass or forb seeds are 

rare or hard to collect or propagate; if species are difficult to 

establish from seeds on disturbed sites; or when the project 

requires restoring threatened or sensitive species (Steinfeld 

and others 2007).

Container seedlings can range in size from tiny liners 

and plugs to large pots containing many gallons of growing medium. Because containers come in many sizes and 

shapes (figure 3.23), they can be matched to the project 

objectives and site conditions. When ordering container 

plants, age, stem diameter, height, root size, and depth 

are usually specified, in addition to species and seed 

source. See Chapter 7, Containers, for more information.

60 Tropical Nursery Manual

Figure 3.22—Container plants (left) and bareroot plants (right) 

have different morphological characteristics. Container plants 

come in many types and sizes but all form their roots into a plug 

resulting in less disturbance to the root system during harvest and 

outplant. Illustration from Dumroese and others (2008).

Figure 3.21—Bareroot coconut plants (A) are grown in fields 

and are harvested and shipped with no soil surrounding the roots. 

Tropical soils are challenging to work with, so bareroot plants are 

usually produced in raised beds (B). Photos by Thomas D. Landis.

B

A
The distinguishing feature of container seedlings is that, 

because the roots are restricted, they bind growing media 

into a cohesive “plug” (figure 3.23) making outplanting 

easier, especially on harsh sites.

Wildings

Plants are typically grown in a nursery. For some projects, however, plants are salvaged from areas, such as development sites or roadsides, before planned disturbances. 

Salvaged plants (sometimes called wildings) can be an 

important component of protecting native plant diversity. 

Sometimes salvaged plants are simply relocated quickly 

from one area to another. At other times, plants may be 

transplanted into a nursery, cared for, and outplanted at a 

later time (Steinfeld and others 2007).

Nonrooted Cuttings

Long “pole” cuttings are a common type of nonrooted 

cutting used extensively in the tropics as live fence posts. 

Sometimes called “quick sticks,” these nonrooted stakes 

of easy-to-root species, such as Gliricidia sepium, Erythrina species, or Bursera simaruba, are cut from the major 

branches or stems of trees. They are inserted into the 

ground with a minimum of 1 ft (30 cm) of the cutting in 

contact with the soil (ideally more). Pole cuttings are usually at least 4.0 ft (1.2 m) long and at least a 2-in (5-cm) 

diameter. Sometimes pole cuttings are used in riparian 

restoration projects (Hoag and Landis 2001). For this 

application, cuttings are often 6.0 ft (1.8 m) or more in 

length and up to 8- to 12-in (20- to 30-cm) in diameter 

so that they can be inserted deep enough such that the 

Defining the Target Plant 61

Figure 3.23—Nursery stock can be grown in many sizes and shapes. Consider the advantages and drawbacks of different options 

when defining the target plant materials for a project. Illustration adapted from Steinfeld and others (2007) by Jim Marin.
butt ends remain in contact with the water table. Pole cuttings are very effective in stabilizing stream or riverbanks 

because they resist erosion. When large numbers of poles 

are required, they can be grown in stooling beds in nurseries to avoid the negative effect of collecting from wild 

“donor” plants.

Rooted Cuttings

It is more effective to root cuttings of most woody species in a nursery before outplanting them on the project 

site. A 2- to 4-in (5- to 10-cm) stem section can be used 

(figure 3.24), but it needs to have a healthy bud near the top 

(Dumroese and others 2003). Some species, such as sweet 

potato (Ipomoea batatas), can be propagated from nonrooted vine cuttings, but may be easier to establish in the 

nursery if they are first allowed to root a little in water or a 

planting media before transplanting into containers. See 

Chapter 10, Vegetative Propagation, for more information about collecting and culturing rooted cuttings.

Rootstock

Rootstock refers to specialized roots, such as bulbs 

and corms, and to modified underground stems, such 

as rhizomes and tubers (figure 3.25). Rootstock can be 

62 Tropical Nursery Manual

Figure 3.25—Rootstock can be used to establish some grasses, 

sedges, forbs, and wetland plants that cannot be direct seeded or 

outplanted as seedlings. Photo by Thomas D. Landis.

Figure 3.24—Rooted cuttings use a shorter section of stem with a bud (A). Cuttings quickly grow into large plants under nursery 

culture (B). Illustration A from Dumroese and others (2008), and photo B by Thomas D. Landis.

A B
used for the vegetative propagation of certain grasses, 

wetland plants, food plants, and even some trees. Examples of species propagated from rootstock include yams 

(Diosorea species), kava (Piper methysticum), and breadfruit (Artocarpus species). Bananas and plantains (Musa

species) are often propagated from their corms or root 

structures.

7. What Are the Best Outplanting 

Tools and Techniques?

Each outplanting site has different climatic and soil 

conditions, so outplanting tools and techniques must be 

matched accordingly. Nursery managers must know in 

advance which planting tools will be used so they can 

develop proper plant material specifications, especially 

root length and volume or cutting length and diameter.

No single tool or technique will work well under all site 

conditions. For example, plants with narrow, long root 

systems may not be the appropriate root shape and size if 

deep, narrow holes are impossible to dig efficiently on the 

site because of rocky or muddy conditions. It is important 

to ensure that the planting holes are large and deep enough 

so that the seedlings can be planted properly.

Hand tools such as shovels (figure 3.26), pick mattocks, planting hoes (“hoedads”), and planting bars are 

very popular for outplanting native plants. Plants grown 

in deeper “tall pots” used in many restoration projects 

may require specialized outplanting equipment. Nursery 

managers must work closely with clients to make certain 

that their target plants can be properly outplanted in the 

soil conditions on the project site. A more complete discussion of outplanting tools and techniques is in Chapter 

17, Outplanting.

8. What Is the Best Time for 

Outplanting?

Each site has an optimal time when chances for plant survival and growth are greatest—the “outplanting window.” 

The outplanting window is usually defined by looking 

at the climate and historical information from Question 

2 and the limiting factors described in Question 3. For 

example, in some tropical areas, soil moisture is the main 

limiting factor. In these cases, the outplanting window 

is at the onset of the rainy season when soil moisture is 

increasing and evapotranspirational losses are low (figure 

3.27) Areas without a dry season may have other limiting factors, such as heavy rainfall and flooding, that define 

outplanting windows differently. The specific dates of outplanting windows will change with latitude and elevation.

Defining the Target Plant 63

Figure 3.26—The type of outplanting tool to be used is critical 

to defining the size and shape of the target plant. Photo by Thomas 

D. Landis.

Figure 3.27—The outplanting window is the period of time in 

which site conditions are the most favorable for plant survival and 

growth. The window will vary among geographical locations. 

For sites with a pronounced dry season, the outplanting window 

occurs at the onset of the rainy season. Illustration adapted by 

from South and Mexal (1984) by Jim Marin.
In producing target plants for a project, the nursery 

must work backwards from the project’s outplanting window to schedule propagation. This approach ensures the 

plants will be ready at the beginning of the outplanting 

window, as described in Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records.

Learning and Adapting: Field 

Testing the Target Plant

Starting small and expanding on successes is a basic principle of effective project management (Mollison and Slay 1991). 

If land managers are able to take a season or more to test out 

some plant materials and strategies before committing on a 

large scale, they can learn important lessons and ultimately 

increase their successes. At the start of any planting project, 

the land manager and the nursery manager need to agree on 

certain morphological and physiological specifications based 

on answers to the eight questions that define a target plant. 

This prototype target plant is grown in the nursery, and its 

suitability is then verified by outplanting trials that monitor 

survival and growth.

Monitoring survival and growth during the first few 

months after outplanting is critical because problems can 

show up soon after planting. Problems with seedling quality, poor planting, or exposure to drought conditions result 

in plants gradually losing vigor and perhaps dying. Therefore, 

plots must be monitored during the first month or two after 

outplanting and again at the end of the first year for initial 

survival. Subsequent checks after 3- and 5-year periods will 

give a good indication of plant growth rates. This performance information is then used to give valuable feedback to 

the nursery manager who can work with the client to refine 

the target plant specifications for the next crop (figure 3.1).

The strategy of starting small is not always possible. 

Some projects are urgent and require full planting as soon 

as possible. Clients with projects that must be planted all at 

once will not be able to benefit from the learning opportunities generated by the target plant feedback cycle. In such 

cases, the best available information and experience is used 

to define the target plants to immediately serve the client’s 

needs. When the project is complete, however, the nursery 

and other people involved can still learn from the outcomes 

and apply the lessons to future projects of a similar nature.

References

Ares, A.; Fownes, J.H.; Sun, W. 2000. Genetic differentiation of 

intrinsic water-use efficiency in the Hawaiian native Acacia koa. 

International Journal of Plant Sciences. 161(2): 909–915.

Bean, T.M.; Smith, S.E.; Karpiscak, M.M. 2004. Intensive revegetation in Arizona’s hot desert: the advantages of container stock. 

Native Plants Journal. 5: 173–180.

Cerro Nara Rainforest Conservation. 2010. Analog forestry 

employed in Cerro Nara. Central West Coast, Costa Rica: ProNara. http://www.cerronara.org/. (August 2010).

Dumroese, R.K.; Wenny, D.L.; Morrison, S.L. 2003. A technique 

for using small cuttings to grow poplars and willows in containers. Native Plants Journal. 4: 137–139.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008 Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Hoag, J.C.; Landis, T.D. 2001. Riparian zone restoration: field 

requirements and nursery opportunities. Native Plants Journal. 

2: 30–35.

Kramer, A.T. 2007. Successful restoration of plant communities: why pollinators matter. Lecture at the Chicago Botanic Garden. http://www.chicagobotanic.org/downloads/staff/kramer/

Kramer_071907DonorTalk.pdf. (August 2010)

Landis, T.D.; Dreesen, D.R.; Dumroese, R.K. 2003. Sex and the 

single Salix: considerations for riparian restoration. Native 

Plants Journal. 4: 110–117.

Landis, T.D.; Lippitt, L.A.; Evans, J.M. 1992. Biodiversity and 

ecosystem management: the role of forest and conservation nurseries. In: Landis, T.D., ed. Proceedings, Western Forest Nursery 

Association. Gen. Tech. Rep. RM-221. Fort. Collins, CO: U.S. 

Department of Agriculture, Forest Service, Rocky Mountain Forest and Range Experiment Station: 1–17.

Landis, T. D. 2011. The Target Plant concept - a history and 

brief overview. In: Riley, L.E., Haase, D.L. and Pinto, J.R. tech 

coords. National Proceedings: Forest and Conservation Nursery 

Associations – 2010. Proceedings RMRS-P-65. Fort Collins, CO: 

US Department of Agriculture, Forest Service, Rocky Mountain 

Research Station, Proceedings. 61-66.

64 Tropical Nursery Manual
Mollison, B.; Slay, R.M. 1991. Introduction to permaculture. 

Tyalgum, Australia: Tagari Publications. 198 p.

South, D.B.; Mexal, J.G. 1984. Growing the “best” seedling for 

reforestation success. Forestry Department Series 12. Auburn, 

AL: Auburn University. 11 p.

St. Clair, B.; Johnson, R. 2003. The structure of genetic variation and implications for the management of seed and planting 

stock. In: Riley, L.E.; Dumroese, R.K.; Landis, T.D., tech. coords. 

National proceedings: forest and conservation nursery associations—2003. Proceedings RMRS-P-33. Ogden, UT: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research 

Station: 64–71.

Steinfeld, D.E.; Riley, S.A.; Wilkinson, K.M.; Landis, T.D.; Riley, 

L.E. 2007. Roadside revegetation: an integrated approach to 

establishing native plants. Vancouver, WA: Western Federal 

Lands Highway Division. 

Sutton, R. 1980. Evaluation of stock after planting. New Zealand 

Journal of Forestry Science 10: 297-299. 

Additional Reading

Guinon, M. 1993. Promoting gene conservation through seed and 

plant procurement. In: Landis, T.D., tech. coord. Proceedings, 

Western Forest Nursery Association. Gen. Tech. Rep.. RM-221. 

Ft. Collins, CO: U.S. Department of Agriculture, Forest Service, 

Rocky Mountain Forest and Range Experiment Station: 38–46.

Jeffrey, J.; Horiuchi, B. 2003. Tree planting at Hakalau National 

Wildlife Refuge—the right tool for the right stocktype. Native 

Plants Journal. 4: 30–31.

Landis, T.D.; Dumroese, R.K.; Haase, D.L. 2010. The container 

tree nursery manual. volume 7: seedling processing, storage, and 

outplanting. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 188 p.

Millar, C.I.; Stephenson, N.L.; Stephens, S.L. 2007. Climate 

change and forests of the future: managing in the face of uncertainty. Ecological Applications. 17(8): 2145–2151.

Defining the Target Plant 65
66
67

A successful nursery provides healthy, high-quality plant materials ready to plant 

when clients need them. For areas with a pronounced dry season, clients usually need 

plants at the beginning of the rainy season. For areas with adequate rainfall all year or 

that can be irrigated, the outplanting date may be anytime. In addition to being ready 

on time, plants must also be of the correct species, quantity, genetic source, size, age, 

and container type, as defined by target plant requirements described in Chapter 3, 

Defining the Target Plant (figure 4.1). To ensure that plants germinate, grow, remain 

healthy, and become hardy to survive outside the nursery, all the plant’s environmental and nutritional requirements must be met while they are in the nursery. These 

requirements change as the plants develop. The planning and scheduling to meet 

these requirements is called nursery crop planning. Crop planning coordinates time, 

resources, labor, and space to produce a healthy crop of plants on time. Throughout 

this chapter, the following example of how target plant requirements work with nursery crop planning is used.

In March, a retired cattle rancher calls your native plant nursery. To leave a legacy for her grandchildren, she wants to plant 500 koa trees (Acacia koa, an endemic 

species important in many Hawaiian forests) on a former cattle pasture now fenced 

and free of cows. Her property is at 2,000-ft [610-m] elevation on the leeward side of 

the Big Island of Hawai‘i. The area has been in pasture for more than a century. She 

had the soils analyzed, and they are typical from a nutrient standpoint. She wants 

easy-to-plant trees. Although the rainy season starts in mid-November, she plans to 

plant with help from her son, daughter-in-law, and two grandchildren during the 

holiday, so her ideal delivery date is December 15. She plans to remove the grass 

from the planting areas and then plant the seedlings using a mattock and shovel.

With these details, you can now begin to work with the client to establish target 

plant requirements for her koa seedlings (figure 4.1).

Crop Planning: Propagation 

Protocols, Schedules, and Records

Kim M. Wilkinson and Douglass F. Jacobs

4

Facing Page: Crop planning organizes schedules, resources, labor, and space to produce a healthy crop of plants on schedule. 

Victoria Henry records some information on a crop in the Agroforestry Research Greenhouse at the University of the Virgin 

Islands, Agriculture Experiment Station in St. Croix. Photo by Brian F. Daley.
1. Objective—Native reforestation/legacy planting with 

locally adapted Acacia koa.

2. Site Evaluation—Client’s soil analysis is unremarkable; she has looked at existing vegetation (pasture 

grasses, weeds) and realizes control is needed; she 

examined climate data for the area, especially historical trends of when the rainy season starts and ends, 

and the average rainfall.

3. Limiting Factors on Outplanting Site—Likely shortage of viable population of appropriate beneficial local 

microorganisms (Rhizobium bacteria and mycorrhizal 

fungi); competing grass and weeds; minor risk of a 

cow or horse eating the young trees.

4. Mitigating Measures for Limiting Factors—Seedlings 

will be inoculated with Rhizobium bacteria and 

mycorrhizal fungi in the nursery. Client will remove 

competing grass and weeds before planting; advise 

her to mulch around trees and diligently continue 

to control weeds. Client has fenced site to minimize 

risk of a cow or horse getting in and eating the seedlings and will keep secure.

5. Genetic Considerations—Seeds sourced according 

to transfer guidelines from forestry department for 

locally adapted, genetically appropriate koa for this 

site. In this case, seeds will be collected (by permission) from a nearby koa forest at a similar elevation: 

minimum 50 parent trees of good form.

6. Type of Plant Materials—500 Acacia koa seedlings 

from genetic sources listed previously. Containers: 

Ray Leach “Stubby” cells; Size: 15 cm height, 3.5 mm 

stem diameter; Roots: firm and nodulating with 

Rhizobium, inoculated with mycorrhizal fungi 

(AMF). Seedlings will be watered thoroughly while 

still in their containers immediately before they are 

outplanted.

7. Outplanting Tool or Technique—Seedlings will be 

outplanted with a mattock and shovel. After outplanting, seedlings will be mulched with a biodegradable 

weed barrier topped with macadamia nut husk mulch, 

being careful not to let the mulch touch the stems. 

Trees will be flagged with a bamboo stake and bright 

flagging for ease of monitoring and maintenance.

8. Timing of Installation/Outplanting Window—This 

area’s rainy season usually begins in November, but 

labor will not be available until December. Target date 

for seedling delivery is December 15.

For complex, large, or specialized projects, the assistance 

of a professional planner is usually needed to assess the site 

and create a plan, including appropriate species selection 

and determination of the other target plant needs for each 

species to be planted. Examples of more specialized plans 

include projects that warrant a Forest Stewardship Plan, 

a Conservation Plan, a Habitat Restoration Plan, Farm 

Plans (such as the U.S. Department of Agriculture, Natural 

Resources Conservation Service might do for soil conservation or riparian areas) and others. If your client requires this 

level of assistance, steer him or her to the appropriate agency 

or professional and invite the client to place an order when 

the plan is ready. See Chapter 18, Working With People, for 

more information on client education and project planning.

Based on the client’s target plant requirements, including the agreed-upon delivery date, the nursery manager 

can now schedule crop production (figure 4.2). Crop production generally includes these activities:

• Developing and refining propagation protocols for the 

species so that requirements are met during germination, establishment, rapid growth, and hardening.

• Developing growing schedules for the crop based on 

the crop’s three growth phases (establishment, rapid 

growth, and hardening) and the distinct requirements 

for each phase.

• Developing facilities schedules to ensure that space, 

labor, equipment, and supplies to support the crop 

(and the other crops being grown simultaneously) 

during all growth stages are available.

• Keeping written records, including a daily log and plant 

development record, so that any shortcomings can be 

corrected and successes can be replicated in future crops.

68 Tropical Nursery Manual

Figure 4.1—Crop planning begins with defining specifications of 

the “target plants” the nursery will 

produce, as described in Chapter 3, 

Defining the Target Plant. Adapted 

from Landis (2011) by Jim Marin.
Developing Propagation Protocols

A propagation protocol describes all the steps necessary 

to grow a species under local conditions, from the collection of seeds or cuttings all the way through shipping the 

plants to the client. It is meant to be a reliable, repeatable 

guide to producing and scheduling a crop of that species. 

It will also help you coordinate the production of all crops 

being grown simultaneously in the nursery.

A protocol is ideally comprehensive, systematic, and 

detailed, much like a cookbook recipe. Protocols with the 

most detailed information make it easier to plan and schedule 

the next crop. The example protocol in table 4.1 shows the 

type of information usually included. The exact schedule and 

performance of any species will vary greatly depending on the 

unique conditions of the nursery and on other variables, such 

as seed sources and weather patterns so protocols may need 

to be adjusted accordingly to produce the best plants.

After protocols are developed, they are refined with each 

crop, leading to dramatic improvements in nursery efficiency 

and effectiveness from season to season. Propagation protocols serve as an essential guide for planning and scheduling 

each crop. 

A protocol typically describes the following aspects and 

characteristics:

• Species name and ecotype.

• Time necessary to grow to target plant specifications.

• Target plant specifications (for example, height, root 

system, and stem diameter).

• Propagule sources, characteristics, collection, and 

processing.

• Preplanting propagule treatments.

• Growing area preparation.

• Management for, and length of, establishment, rapid 

growth, and hardening phases.

• Harvesting and shipping practices.

• Outplanting window and planting technique(s).

Draft a Protocol

The protocol is developed using firsthand experiences 

and outside sources of information. If little to nothing is 

known about the species, the process of drafting a protocol 

can help you organize what is known and take an educated 

guess at how to proceed with growing a particular species. 

Start by systematically searching relevant literature published by trade journals, native plant societies, and botanical 

gardens. An excellent source of propagation protocols, continually updated by growers and free of charge, is online at 

the Native Plant Network (http://www.nativeplantnetwork.

org) (Landis and Dumroese 2000; 2002). If specific information on the desired species is unavailable, try to find another 

species within the same genera or even a related species 

grown in similar climatic zones, to see if any information may 

be applicable. Next, gather information from observations 

of how the plant grows in nature. This information may be 

observed first hand in the field and by asking local people who 

are familiar with the plant. Although collectors of plant materials for cultural uses may have never propagated the species, 

Crop Planning: Propagation Protocols, Schedules, and Records 69

Figure 4.2—The diversity of species grown in native plant 

nurseries (A) calls for detailed crop planning and scheduling to 

ensure that high-quality plants (B) are delivered to clients when 

they need them. Photo A by Diane L. Haase, and photo B by 

Douglass F. Jacobs. 

B

A
they are likely to know the plant’s life cycle such as when and 

how seeds are dispersed. Finally, seek advice from other nursery managers. Although some managers of private nurseries 

may be disinclined to share their propagation methods, government nurseries and botanical gardens are often excellent 

sources of information.

Based on the information gathered, the first draft is the 

best-informed guess of what will be required to grow a species in your nursery the first time. The plants themselves 

will prove the protocol right or wrong as they grow. Nursery 

records, including the daily log and plant development record 

described later in this chapter, will enable a comparison 

between projected development and actual growth.

Test and Adjust Protocols

Refine the protocol with site-specific information from 

your nursery after the production of each crop. Do not be 

discouraged if a protocol drafted from background research 

or another nursery’s experience does not produce the same 

results; the goal is to adjust the protocol to reflect your nursery 

conditions. Year-to-year variations in weather or unforeseen 

operational changes often keep crops from growing exactly as 

projected. Allow room for flexibility and make adjustments 

based on observed factors. You may want to tinker with growing media ingredients, seed germination methods, irrigation 

practices, and so on. Keeping records is key. Sometimes 

new information and discoveries will significantly improve 

70 Tropical Nursery Manual

Family scientific name Fabaceae

Family common name Legume

Scientific name Acacia koa Gray

Common name Koa

General 

distribution

Native to six major Hawaiian Islands: Hawai‘i, Moloka‘i, Maui, Lana‘i, O‘ahu, and Kaua‘i. Original range: 

300–7,000 ft (90–2,134 m). Today, most thriving koa trees are found between 3,000 and 6,000 ft (915 and 1,830 

m) elevation. Introduced pests and diseases limit their presence below 2,000 ft.

Propagation goal Plants (tree seedlings)

Propagation method Seed

Product type Container: 115 ml Ray Leach “Stubby” Cell

Time to grow 16 to 18 weeks

Target specifications Seeds from locally adapted, genetically appropriate source matched to outplanting site; seedlings approx. 15 cm 

tall; diameter 3.5 mm; and roots firm and nodulating with Rhizobium.

Propagule collection Genetic quality is crucial; source must be carefully matched with the outplanting site before seeds are collected. 

Diversity is also important to withstand diseases; collect at least 50 mother trees, throughout the canopy. Pods are 

about 6 in (15 cm) long and 1 to 1.5 in (2 to 4 cm) wide, with 6 to 12 seeds per pod. Pods are ready to pick when 

brown, and when opened the seeds inside are deep brown and full (not green, flat, or small). Seeds can also be 

collected from the ground. In some populations, koa seeds can be collected any time of year, in other populations 

August to October (end of dry season) is best time to collect.

Propagule processing Pods are dried in the sun until they can be opened easily. Seeds are extracted by hand or by threshing. Once out 

of the pods, seeds are dried more as necessary (ideal moisture content 6 to 8%). Dried seeds can be stored in an 

airtight container away from direct sunlight. Properly dried seeds can store for 12 to 24 months at room temperature, many years longer in cooler conditions.

Seeds/kg Seed size is highly variable, a kilo of processed seed can contain between 5,000 and 15,000 seeds (2,500 to 7,500 

seeds/lb).

Purity 100%.

Germination 70 to 90% (can be lower depending on weather conditions during ripening).

Pre-sowing 

treatments

Scarification is required. Mechanical scarification (nicking with a nail clippers on the side opposite the point of 

attachment to the pod) is used for small lots. Hot water treatment (195 °F, 90 °C) in a volume ratio of at least 5 

parts water to 1 part seeds for 1 to 3 minutes. In either case, scarified seeds are soaked overnight to allow water to 

penetrate into the seed. Seeds germinate in 2 to 7 days.

Growing area If possible, some cover (greenhouse or temporary cover) is ideal for the first 2 weeks after germination to protect 

sprouts from seed-eating birds and rodents and from hard rains. Thereafter, uncovered growing areas work well.

Table 4.1—An example protocol for Acacia koa, adapted from Wilkinson and Elevitch (2003; 2004). 
Crop Planning: Propagation Protocols, Schedules, and Records 71

Growing medium Most well-drained media work fine. One example media is 50% peat moss, 25% perlite, 25% vermiculite, amended 

with compost, dolomite lime, gypsum, and triple super phosphate. Media should also be inoculated mycorrhizal 

fungi, local strains if available. Seedlings will be inoculated with Rhizobium at 2 weeks.

Establishment phase Scarified seeds will germinate in 2 to 7 days. They may be direct seeded after scarification (1 seed/cell) or, for less 

viable seedlots, seeds (sown on paper towels or in beds) are transplanted into tube containers that have been prefilled with medium (1 seed/cell) as soon as the root begins growing from the seed. Cover with growing medium 

shallowly (about 1/4 inch or 0.6 cm deep), followed by a thin mulch layer such as #2 poultry grit. Water with a 

fine-headed sprayer to keep moist, but not wet. Full sun is best. Daily water is usually necessary, by hand or with 

an automated system. Protect from bird and rodent predators who are attracted to the sprouts. At 2 weeks of age, 

inoculate with Rhizobium using a slurry made from native nodules collected from seed source areas.

Length of 

establishment phase 2 to 3 weeks

Rapid growth phase Seedlings are watered daily, usually in the morning. Especially hot, dry days may necessitate a second watering 

in the early afternoon. (Late afternoon and evening watering is not recommended, as it facilitates pest problems, 

such as sooty molds.) The media should not be allowed to dry out. After about 4 weeks, check to ensure Rhizobium

inoculation was effective. After about 6 weeks, seedlings may be double-spaced in the Ray Leach trays (from 98 

trees per tray to 49 trees per tray) to ensure each seedling receives full sunlight and good air movement to facilitate 

strong stem development. At the period, depending on seed lot, about 5 to 10% of the seedlings will be apparent 

poor performers, and these should be culled. Remaining seedlings are monitored for pests, but pests are generally 

not problematic in the nursery. If any weeds enter the media, they should be removed. No fertilizer application as 

necessary if seedlings were inoculated with Rhizobium and mycorrhizal fungi. No pruning.

Rapid growth phase 7 to 10 weeks

Hardening phase Seedlings should never be allowed to dry out, but watering frequency is reduced to introduce seedlings to temporary, moderate water stress. If the outplanting site is an especially windy area, “brush” seedlings gently using a 

length of PVC pipe to improve stem strength. Seedlings remain outside, exposed to full sunlight, not fertilized.

Length of 

hardening phase 6 weeks

Harvesting, storage,

and shipping

When seedlings have reached target size, they may be delivered to the planting site. Water thoroughly before shipping and just before outplanting. They are not extracted from their container until the moment they are planted, as 

keeping them in the container is necessary to protect the roots and the viability of the Rhizobium nodules. Seedlings 

may be stood up in buckets, cardboard boxes, or delivered in their trays. Seedlings must be protected from wind and 

excessive heat during transport, but refrigeration is not recommended. Empty containers and trays may be returned 

after the planting is complete.

Outplanting and

cultivation comments

Planting areas must be fenced to exclude any grazing animals, and competitive vegetation must be removed. At 

planting, seedlings should be carefully removed from their container and planted at the correct depth, so the 

ground is even with the root collar. Soil should be firmed around the tree, muddied in with water if possible. A weed 

barrier/mulch around the tree (but not touching the trunk) can aid establishment.

Citations Wilkinson, K.M.; Elevitch, C.R. 2004. Propagation protocol for production of container Acacia koa Gray plants; 

permanent agriculture resources, Holualoa, Hawai‘i. In: Native Plant Network. URL: http://www.nativeplantnetwork.org 

(September 2013). Moscow, ID: University of Idaho, College of Natural Resources, Forest Research Nursery.

Table 4.1 (continued)
production methods (see Chapter 20, Discovering Ways to 

Improve Nursery Practices and Plant Quality) and will be 

added to the protocol. As the protocol is updated, the nursery 

develops an increasingly accurate and helpful guide for how 

to grow each species.

Developing Growing Schedules

With the propagation protocol in hand, it is time to 

develop a growing schedule that covers all phases of crop 

production and the time necessary to complete each step. 

When the timing for nursery crops is understood, appropriate dates for sowing seeds or striking cuttings can be 

calculated by counting backwards from the desired outplanting date. Knowing when propagules can be collected 

and how long the species will take to produce enables the 

nursery to work with clients’ schedules. For example, koa 

requires 18 weeks to grow, including time for preparing 

seeds for sowing. So, if plants are needed for outplanting 

on December 15, seeds must be ready for sowing by August 

11. The total time required for the production of each crop 

will vary by species, container size, season, and nursery 

environment. Genetics and the variability of seedlots may 

also cause variations in crop scheduling even for the same 

species. Less well-known or more temperamental species 

require the nursery manager to build a safety margin of 

2 to 4 weeks into the schedule in case of problems with 

germination or growth. The growing schedule focuses on 

these steps as plants move through the nursery: propagule 

collection, cleaning, and treating; the three phases of plant 

growth (establishment, rapid growth, and hardening); and 

storage and shipping. Although other chapters cover these 

topics in detail, in this chapter, they will be discussed in 

general terms as they relate to scheduling.

Propagule Collection, Processing, 

and Treatment

Collection

Most often, nurseries growing native and culturally 

important plants cannot obtain their seeds or cuttings 

from a central supplier or mail-order catalog. Therefore, 

key questions are where, when, and how to collect seeds or 

cuttings. The where is usually answered by the location of 

the outplanting sites: locally adapted, genetically appropriate materials should be collected from the same or similar 

areas as the outplanting site (figure 4.3A). For some native 

species, usually commercially important trees, recommendations for appropriate sources or seed zones may be available, as mentioned in Chapter 3, Defining the Target Plant. 

For most other native species, the nursery will have to do 

its best to research, define, and obtain locally adapted and 

genetically appropriate propagules for its client’s site. The 

when question is answered by the plants living in the collection sites: when does this species, in the appropriate 

collection site(s), produce seeds? (Or for vegetative propagation, when is the best time to collect cuttings?) For 

some species, it may be possible to collect throughout the 

year. For other species, a narrow window may be all that 

is available, and if this window is missed, orders will have 

to wait until the following year. Seeds of some species may 

store well for many years, enabling the nursery to develop 

collections to keep on hand. Finally, how to collect may 

be determined based on published literature, local knowledge, and experience. To provide plants when the client 

needs them, the nursery manager has to account for the 

time needed to obtain the propagules and grow the plants.

Processing

Propagules need to be processed immediately after collection. For most seeds, processing helps ensure that the 

72 Tropical Nursery Manual

Figure 4.3—Nursery crop planning includes scheduling propagule 

collection (A), processing, and seed treatments (B) to produce target 

plants by the delivery date. Photo A by J.B. Friday, and photo B 

by Craig R. Elevitch.

B

A
seeds stay dormant until they are needed. Depending on 

the species, seeds may need to be processed in one or more 

of the following ways: extraction from pods or fruits, washing, winnowing, sorting, grading, and drying. Some seeds 

do not store and must be sown when they are fresh. Cuttings may need to be dried, soaked, placed in rooting beds, 

or other treatments. Processing requirements and the time 

and space needed to complete them need to be considered 

as part of the schedule.

Treatment

Seeds of native species, and sometimes seedlots of the 

same species, vary widely in dormancy, so seed treatments need to be scheduled properly. Similarly, the optimum time to harvest cuttings varies among native species. Scheduling when to treat propagules is important 

for planning a target sowing or cutting establishment 

date for the crop (figure 4.3B).

Crop Growth Phases

Because a tiny germinant has different needs than a 

large plant that is nearly ready for outplanting, understanding the growth phases of crops is essential to nursery planning. The development of most nursery crops 

can be divided into three phases: establishment, rapid 

growth, and hardening (figures 4.4, 4.5). Plants in each 

phase have distinct requirements for light, water, nursery 

space, and the types of attention and labor necessary to 

keep them healthy. The nursery manager’s objectives for 

the crop are also different at each phase. Table 4.2 summarizes some typical aspects of each of the three phases.

Establishment

For plants grown from seeds, the establishment 

phase is defined as the time from the sowing of the seeds 

through the germination, emergence, and development 

of the first true leaves or primary needles (figure 4.6A). 

For plants grown from cuttings, the establishment phase 

extends from placing cuttings into containers through 

the initial development of roots and shoots. This phase 

is of critical importance because mistakes can prevent or 

delay emergence or rooting. For seeds, some species require 

light to germinate and need to be surface sown while seeds 

Crop Planning: Propagation Protocols, Schedules, and Records 73

Figure 4.4—Understanding the growth phases that crops go through is essential to crop planning. The development of most nursery crops 

can be divided into three phases: establishment, rapid growth, and hardening. Adapted from Dumroese and others (2008) by Jim Marin.

Figure 4.5—Changes in seedling morphology during the three 

growth phases. Growth is relatively slow during the establishment phase. Most shoot growth occurs during the rapid growth 

phase, which ends when target height specifications are met. During hardening, roots continue to grow, resulting in an increase in 

seedling stem diameter (caliper). 
of other species may be covered with mulch. For cuttings, 

some species need rooting hormones applied at the proper 

rates and timing. The establishment phase typically lasts 4 

to 8 weeks, although species slow to germinate from seeds 

or root from cuttings may take 1 year or more. The main 

goal of this phase is to maximize survival with uniform germination and establishment of the plants.

Rapid Growth

During the rapid growth phase, plant shoots increase 

dramatically in size, often approaching target size (figures 

4.6 B and C). Plants need to be somewhat protected during this phase to encourage rapid (but not excessive) shoot 

growth and minimize stress.

Hardening

Although it is relatively easy to grow a seedling to target size, the tricky part is the hardening phase. During 

hardening, energy is diverted from shoot growth to root 

growth (figures 4.6 D, 4.6 E). Hardened plants are conditioned to endure the stresses of shipping, handling, and 

outplanting and fortified so that they have the energy and 

nutritional reserves to survive and grow after outplanting. If the hardening phase is too short and plants do not 

74 Tropical Nursery Manual

Table 4.2—The three general phases of crop development for seedlings. After the three phases of crop development are understood for a 

species, the growing and facilities schedules can be developed to meet crop needs during each phase. Adapted from Landis and others (1998).

Phase Establishment Rapid growth Hardening

Definition From germination through 

emergence and formation of true 

leaves.

From emergence of true leaves to when 

seedling approaches target size. Rapid 

increase in size, particularly in terminal 

shoot.

Energy diverted from shoot to root growth; 

seedling reaches target height and stem diameter; seedling is conditioned to endure stress.

Duration Typically 14 to 21 days for germination; 4 to 8 weeks for early 

growth.

Varies widely, typically about 

10 to 20 weeks.

Varies widely, normally from 

4 to 12 weeks.

Objectives • Fill containers efficiently

• Maximize uniform 

germination

• Maximize survival

• Minimize damping-off

• Minimize stress

• Encourage shoot growth

• Maintain environmental factors 

near optimum levels

• Monitor as seedling approaches 

target height and roots fully 

occupy container

• Slow shoot growth

• Encourage root and stem 

diameter growth

• Acclimate to outplanting environment

• Condition to endure stress

• Fortify for survival after outplanting

Special

needs

• Protect from weather and 

pests

• Keep temperature warm

• Irrigate to keep soil 

“moist, but not wet”

• No or low fertilizer

• Protect from stress

• Monitor sun exposure 

• Irrigate appropriately

• Fertilize properly

• Induce moderate moisture stress

• Progressively expose to sun equivalent 

to outplanting conditions (full sun or 

partial shade)

• Expose to ambient temperatures 

and humidity

• Provide good air flow/wind

• Reduce fertilization rates and change 

mineral nutrient ratios

Labor

• Monitor germination

• Introduce beneficial microorganisms

• Thin

• Resow and transplant if 

necessary

• Scout for pests and diseases

• Monitor environment

• Modify crop density to encourage 

good development

• Adjust culture to avoid excessive 

shoot height 

• Scout for pests and diseases

• Monitor crops and environment carefully; see Chapter 14, Problem Prevention and Holistic Pest Management, and 

Chapter 19, Nursery Management, for 

details 

• Deliver crops to clients in a timely fashion 

to avoid problems with holdover stock 

• Scout for pests and diseases
Crop Planning: Propagation Protocols, Schedules, and Records 75

have time to reach the appropriate physiological condition, 

plants may still have the correct physical size, but survival 

and growth after outplanting are compromised. Therefore, 

good crop planning ensures adequate hardening before ontime delivery to the client. See Chapter 15, Hardening, for 

more discussion on this topic.

Harvesting and Shipping

After plants are hardened, prompt outplanting is essential to ensure that they can take full advantage of their hardened condition. Proper crop scheduling ensures that a crop 

goes through the three phases of growth and that healthy 

plants are ready for shipping (figure 4.6 F) and outplanting 

on the agreed-upon delivery date. See Chapter 16, Harvesting and Shipping, for more discussion on this topic.

Problems With Holdover Stock

A common problem is the failure of clients to pick up 

plants on schedule. This problem can be avoided by good 

scheduling practices and frequent communication with 

clients, especially by providing periodic updates to advise 

them when seedlings will be ready. In some cases, having 

penalties, such as storage fees, in the contract for late pickups may also encourage clients to pick up their plants in a 

timely fashion. When communicating with clients, emphasize upfront that prompt outplanting is in everyone’s best 

interest, not only for the nursery and the health of the 

plants, but also for the success of the client’s project. See 

Chapter 19, Nursery Management, for more information 

about contracts and communicating with clients.

If the schedule to outplant after hardening is not met, 

problems can develop. When plants are held too long in 

the nursery, the root system becomes woody and loses 

its ability to take up water and nutrients (figure 4.7A). 

Structural problems may also occur; roots may spiral 

(figure 4.7B) and, instead of expanding outward and 

downward into the soil after outplanting, girdle the plant 

or cause it to be unstable in high winds. Shoot growth 

may resume and negatively affect the root-to-shoot ratio 

(figure 4.7C), and the plant loses its resistance to stress. 

Making a growing schedule as shown in table 4.3 is very 

helpful to provide plants on time; neither ready too early 

or too late.

Developing Facilities Schedules

Ensuring that the nursery facilities and resources required 

to meet plant needs through the growing cycle are available at 

the appropriate times is the goal of facilities scheduling. The 

Figure 4.6—Shown is koa (Acacia koa) 

at different growth phases: germination and 

establishment (A), establishment phase 2 

weeks after germination (B), early rapid 

growth phase (C), late rapid growth phase 

(D), hardening phase in outdoor area (E), 

and target plant ready to ship to client (F). 

Photos by Douglass F. Jacobs.

B

C E

F

A

D
76 Tropical Nursery Manual

space, labor, equipment, and supplies required for each crop 

during the different stages of propagation must be planned.

Except for nurseries with elaborate climate control 

systems, crops are often moved from one structure to 

another as they progress through the three development 

phases. See Chapter 5, Propagation Environments, for 

additional information. Using the koa example, crops are 

protected in a special germination area during the establishment phase and then moved to an outdoor growth 

area for rapid growth and hardening (table 4.4). Needs 

differ for various crops and nurseries; for example, 

another species might have seeds in trays on benches, 

then be moved to containers in a shade house, and finally 

transplanted to large pots in full sun for hardening. The 

amount of space the crop will require varies by growth 

phase: emerging seedlings may take up little room, but 

plants take up much more space after they have been 

transplanted into larger containers or spaced more 

widely as they grow larger. Although the example in table 

4.4 does not go into such detail, the facilities schedule 

should include calculations for how much space each 

crop will use, how many hours of labor will be needed, 

and the quantities of materials (such as growing media) 

required during crop production.

A good facilities plan considers crop layout, that is, 

what crops and stocktypes go where in the nursery. This 

layout is planned to effectively provide appropriate growing requirements (temperatures, sunlight) and other cultural requirements (frequency of fertilization, watering, 

or other treatments) for all the crops grown each season. 

Fast-growing species with similar growing and cultural 

requirements usually can be grouped together in one 

area, and moderate and slower growing species can be 

grouped together in another area of the nursery. This 

grouping method allows for species with similar requirements and growth rates to be treated effectively and efficiently. Likewise, the flow of plants out of a protected 

area to an outdoor nursery needs to be taken into consideration so that plants that finish in the same timeframe 

can be moved out and a new crop, if scheduled, can be 

planned for the available empty space.

Facilities scheduling is indispensable in determining 

how resources within a nursery can be best distributed to 

maximize production and minimize conflicts associated 

with overlapping needs (figure 4.8). The facilities schedule 

(table 4.4) may be combined with or posted side by side 

with the growing schedule (table 4.3) and the staff needs to 

have easy reference to it.

Figure 4.7—Crops that are held too long (holdover stock) will 

not be properly conditioned to endure the stresses of transportation 

and outplanting (A). The root systems spiral and the plants become 

rootbound, such as this māmane (Sophora chrysophylla) (B). The 

shoots grow too tall for their roots to support and become top heavy 

(C). Illustration A from Dumroese and others (2008) by Jim Marin, 

photo B by J.B. Friday, and photo C by Thomas D. Landis.

B

C

A
Scheduling Multiple Crops

Most native plant nurseries must deliver a suite 

of diverse species on a single shipping date to meet 

client needs. Therefore, the growing and facilities 

schedules are essential to coordinate production 

of multiple crops of different species. The schedules must reflect the growth rates (time required to 

grow the crop) of each species. For example, without proper scheduling, the faster growing species 

may become overgrown before the delivery date 

or the slower growing species may not be ready in 

time. Therefore, slow-growing species need to be 

sown earlier than fast-growing species so they will 

all be ready to plant at the correct time.

In the example in figure 4.8, the nursery is growing 

six species that need to be ready for the same outplanting date in December. To meet the target outplanting, 

koa needs to be sown in August, as they take about 18 

weeks to grow, but if a client also wanted native sandalwood trees in 1-gallon pots at the same time as 

the koa, nursery work would need to begin 1 year in 

advance to bring this slower growing species through 

Crop Planning: Propagation Protocols, Schedules, and Records 77

Activity Seed collection 

and processing

Seed

treatments

Establishment

phase

Rapid growth

phase

Hardening

phase

Duration 2 days 2 days 3 weeks 10 weeks 6 weeks

Dates Scope July 15; collect 

Aug. 1; process Aug. 2

Aug. 3 to scarify; 

Aug. 4 to sow

Aug. 4 to Aug. 25 Aug. 26 to Nov. 3 Nov. 3 to Dec. 14

Propagation 

environment

Field-collection sites Indoors Germinant area protected from 

slugs, birds, rodents, rain, etc.

Main outdoor growth 

area and full sun

Main outdoor growth 

area and full sun

Fertilization None None In growing media: triple super 

phosphate, dolomite lime, 

gypsum, and mycorrhizal 

fungi; inoculate at 2 weeks with 

Rhizobium

None None

Irrigation None None Daily gentle hand watering to 

keep moist

Once daily by hand to 

saturation

Gradual reduction

Target size at 

end of phase

600 viable seeds plus 

extras to store for 

future orders

600 germinants Not applicable (they will usually be about 1 to 2 in (2 to 5 

cm) tall, but no target is set)

Approx. 10- to 12-cm 

height, 2.5- to 3-mm 

root-collar diameter

15 cm tall, stem 

diameter 3.5 mm; 

Roots: firm with 

Rhizobium nodules

Actions Field collection of 

pods. Extract, dry and 

clean seeds at nursery

Mechanical 

scarification; 

soak overnight in 

clean water

Make potting media; sow 

seeds; inoculate seedlings with 

Rhizobium at 2 weeks of age

Pest management; 

check nodulation 

with Rhizobium; 

space double in trays; 

cull poor performers

If to be outplanted in 

windy areas, “brush” 

daily to simulate 

wind and improve 

stem strength

Table 4.3—This example crop schedule shows the necessary steps in each crop development phase and the time required to complete 

each. This schedule should be posted in the nursery so that staff can track the crop’s development and understand what cultural practices 

are required. If appropriate, the schedule can also be shown to clients to help them fully understand the time required to produce their crop.

Figure 4.8—In this hypothetical schedule from Wilkinson and Elevitch 

(2004), six Hawaiian species must be ready at the same time in December. 

Different stock types and species require more or less time to grow in the 

nursery, depending on many factors. Please note this schedule is hypothetical 

only—timing will vary by location, species, and seed sources. Illustration by 

Jim Marin.
all the phases of development. Some species may be fast growing, such as the native wili-wili, but seed collection sites may be 

seasonal and require more advanced notice to collect. (Please 

note: these examples are hypothetical—timing, especially of 

the seed collection, will vary by location and species.)

Keeping Written Records Makes 

Planning Easier

Propagation protocols, growing schedules, and facilities 

schedules can all be improved each season by keeping good 

records. It is important to keep records of how to replicate 

each crop. How long did it take to produce the crop? What 

materials were purchased? How was the crop fertilized, 

watered, and managed during each growth phase? What did 

we learn that will help us next time we grow this species?

Propagation protocols should be revised on a seasonal or 

annual basis, becoming more accurate each year, which then 

improves the usefulness of the crop and facilities schedules. 

The best way to improve the accuracy of your protocols is 

to keep two kinds of written records:

1. The daily log is a journal that notes nursery conditions, activities, and management practices on a daily 

basis. In other words, what was done?

2. Regular plant growth records describe the development of each crop. In other words, how did the crop 

respond to particular management practices?

The daily log and plant development records interrelate 

with the propagation protocols (figure 4.9).

Daily Log

A daily log is a key record that provides a history of nursery management, problem-solving, and crop development 

(figure 4.10). A daily log example is in table 4.5, and appendix 4.B includes a blank daily log form that a small nursery 

might use. Make it a habit to at least jot down something 

each day. Large nurseries may keep more complicated daily 

78 Tropical Nursery Manual

Activity Seed collection 

and processing

Seed

treatments

Establishment

phase

Rapid growth

phase

Hardening

phase

Length 2 days 2 days 3 weeks 10 weeks 6 weeks

Dates Scope July 15; collect Aug. 

1; process Aug. 2

Aug. 3 to scarify; 

Aug. 4 to sow

Aug. 4 to Aug. 25 Aug. 26 to Nov. 3 Nov. 3 to Dec. 14

Labor Two staff members to scout 

out collection site in July 

(1/2 day). Two staff members to go to collection site 

on Aug. 1 and collect (1 full 

day per person). Two staff 

members to dry, process, 

and clean seeds on Aug. 2 

(a few hours intermittently 

throughout day as pods 

dry).

One staff member to 

hand-scarify seeds 

late in day on Aug. 

3, and put them 

in water to soak 

overnight.

Make growing media; fill 

containers; sow seeds; 

hand water daily; monitor 

germination; 

protect from slugs, birds, 

and rodents. Collect 

Rhizobium nodules and 

inoculate seedlings on 

Aug. 18. Update 

client on progress.

Move to outdoor growth 

area, irrigate, monitor 

growth, double-space in 

trays as seedlings grow 

larger, and manage weeds/

pests. Update client as 

end of phase nears. If 

overstock seedlings will 

be available, offer them 

to the client or look for 

another home for them.

Monitor growth;

monitor and 

gradually reduce 

irrigation; brush 

to encourage stem 

diameter. Keep in 

regular touch with 

client. Schedule 

pick-up day and 

time.

Facility/

space 

needed

Permissions and/or permits 

to collect at collection 

site(s). Sunny, level area at 

nursery to lay out pods for 

drying.

Indoors (home or 

nursery office).

Benches in 

germinant area 

(protected).

Benches in main outdoor 

growth area.

Benches in main 

outdoor growth 

area.

Materials

needed

For collection: vehicle, 

pole pruner, pruning ladder, 

collection bags, maps, 

and written collection 

permissions. For processing: tarp and seed storage 

containers.

Seeds, nail clippers, 

a clean container, 

and clean water for 

soaking the seeds.

Scarified seeds,

containers and trays, 

growing media and 

amendments, mycorrhizal 

inoculant, fine-headed 

sprayer and hose for irrigation, blender and nodules to make Rhizobium

inoculant (on Aug. 18).

Extra trays for 

double-spacing 

seedlings, irrigation 

supplies.

PVC pipe or 

bamboo pole for 

brushing, irrigation supplies, 

boxes or buckets 

for transport 

unless plants will 

be transported in 

trays.

Table 4.4. An example of a facilities schedule for Acacia koa.
records and may have separate logs for irrigation, fertilization, 

and the like. Tailor the daily log to suit your nursery. What is 

recorded in the daily log about management practices, environmental conditions, and general crop performance will 

become an invaluable resource for many years to come.

Plant Development Record

Keeping a simple plant development record (or register) for each crop is a great way to build a foundation for 

accurate, site- and species-specific protocols. Some growers choose to record a large amount of detail in their daily 

log and then later summarize information about each crop 

into a plant development record. Others jot notes into a 

plant development record for each crop on a regular basis; 

weekly, monthly, or when the crop is entering a new stage 

in its development.

A plant development record notes what is happening 

with a crop of plants from crop initiation through delivery. 

At a minimum, you can simply put a couple of fresh sheets 

of paper in a three-ring binder, make a tab with the species 

name and date for that crop, and jot down notes on a regular basis as the crop progresses. One way to make it easier 

to keep track of this valuable information is to use a form 

such as the one provided in appendix 4.C or by entering 

information into a computer. An example Plant Development Record is in table 4.6.

Crop Planning: Propagation Protocols, Schedules, and Records 79

Figure 4.9—The three basic crop 

records and how they relate. Use a 

protocol to plan and schedule daily 

activities. Record activities in the 

daily log. The log helps with the 

collection of information about the 

development of each crop. This information is recorded in the plant development record. The protocol is then 

refined and expanded based on this 

new information, which will improve 

production practices the next time 

the crop is grown. Illustrationfrom 

Dumroese and others (2008) by Jim 

Marin.

Figure 4.10—Jot down a few notes every day about what was done 

and what happened with the crop. These records become invaluable 

resources when adjusting protocols and fine-tuning crop and facilities 

schedules. Photo by Tara Luna.
80 Tropical Nursery Manual

Daily log

Date April 4, 2015

Environmental conditions in growing areas 

(light, temperature, and humidity)

Outdoor growing area: 65 °F min; 78 °F max

Sunrise/Sunset times Sunrise: 5:40 a.m. Sunset: 6:55 p.m.

Moon phase Full moon.

Other weather notes 

(cloud cover, and so on)

Partly cloudy, no rain today.

What water did plants receive? 

(irrigation type and frequency or precipitation)

First thing in the morning, automatic overhead watering for 1 hour on most 

plants in main rapid-growth outdoor area. Hand-watered benches 1- to 6-in 

outdoor growing area. Handwatered germination area with fine-headed sprayer 

in morning.

Today’s activities 

(note how many person-hours per activity) 

(fertilization, pest management, transplanting, packing and 

shipping, making potting media, 

moving crops from one structure to another, 

treating or sowing seeds, and so on)

Growing media and filled trays for the new order for 300 seedlings, 2 hours total. 

Handwatered the germinant area and hardening areas, 1 hour. Fixed the small leak 

(noticed yesterday) in the main water line, 1 hour. Answered e-mail correspondence, 1 hour. Fertilized the beach cordia, sandalwood, and ‘ōhi‘a with 200 parts 

per million (ppm) Peter’s 20-20-20 (40 ppm nitrogen), 1 hour. Checked on status of 

milo germinants, 30 minutes. Weed control around perimeter of outdoor growing 

area, 1 hour.

Growth phase status 

(make notes when a crop moves from

one phase to another)

The milo seedlings sown earlier this month are germinating well. I better make 

space in the rapid growth area because I will be able to move them in a couple 

weeks. The sandalwood seedlings that are to be outplanted this December are all 

transplanted and starting the rapid growth phase.

Purchases 

(what supplies or equipment were purchased

and their cost)

Bought a new coupling for fixing the irrigation line plus an extra one to have on 

hand in case of another leak: $10.87. Bought potting media materials for upcoming 

order: $28.45.

Orders 

(what plant materials were delivered and

payments made)

Next week is the pickup of 250 sandalwood seedlings for the high school’s graduating class to plant as a class gift to the community forest. The seedlings are ready; I 

just have to keep the slugs at bay.

General crop/nursery observations Things look good in general.

Questions or concerns There seem to be lots of slugs out... I would like to explore some other organic slug 

control options... maybe I will call around and see if anyone I know has had success 

with that copper barrier stuff and what the cost might be. I need to follow up with 

the Smiths to see if they are ready to confirm koa seedling order for November 

planting—if so I need to give them their contract, collect a deposit, and get planting 

so we can be ready when the rainy season starts.

Table 4.5—An example of a daily log.
Crop Planning: Propagation Protocols, Schedules, and Records 81

Plant development record 

(Koa crop for Waimea Ranch due Dec. 15)

Species name Acacia koa

Propagule source Collection site ABDA, at 2,000 ft (610 m) elevation on the leeward side of the Big Island of 

Hawai‘i from 55 parent trees of good form. 

Date(s) of propagule collection July 15 and Aug. 1

Establishment

Type and length of propagule treatment 

(for example, scarified, stratified)

Hand scarified then soaked in water overnight.

Date of propagule establishment Aug. 2 to 3 

Growing media and tray or container 

type used

Direct sown in Ray Leach Stubby cells; growing media consisting of 50% peat, 25% perlite, 

and 25% vermiculite with amendments and mycorrhizal fungi. 

Germination notes (including date begins and 

ends, percent germination)

Fairly uniform germination of about 92% from Aug. 5 to 12. 

Cutting notes (for example, special 

conditions, hormone treatments)

N/A*

Date transplanted (if not direct sown) N/A*

Container type and growing media 

for transplanting

N/A*

Microorganisms used? Mycorrhizal fungi inoculant in media; inoculated with Rhizobium on Aug. 20, 2014

Misting/Irrigation (type and frequency) Daily hand-watering

Fertilization type, rate, and frequency, if any N/A

Environmental conditions for crop 

(light, temperature, humidity)

Under plastic cover of screened-in greenhouse during establishment so birds and rodents do 

not eat the seeds. 

Horticultural treatments (for example, 

cultivation practices)

N/A*

Date establishment phase completed Aug. 26

Notes (resowing or thinning activities, problems, 

or challenges) 

N/A*

Rapid growth

Time after sowing and sticking to enter rapid 

growth phase 

3 weeks (seeds scarified Aug. 1 and moved to rapid growth area Aug. 25)

Plant size at start of phase (height) About 5 cm tall

Irrigation (type and frequency; 

for example daily, every other day, etc.)

Daily hand-watering

Fertilization type, rate, and frequency N/A*

Environmental conditions for crop 

(light, temperature, and humidity)

Moved to full sun Aug. 27 

Horticultural treatments (for example, 

spacing, cultivation practices)

Spaced out 50% on Sep. 9 

Date rapid growth phase completed Nov. 3, 2014

Notes (development, vigor, and health, 

challenges or problems)

Culled about 5% that were not growing quickly before beginning hardening phase on Nov. 3.

Table 4.6—An example of a plant development record for an Acacia koa crop.
Keep a separate plant development record for each crop 

grown, even if it is just a small trial of a few plants or even if 

the species has been grown before. Any intended improvements, trials, or experiments done with a species (such as 

increasing ventilation in the nursery area or switching to 

a new kind of fertilizer) can also be recorded in a plant 

development record. When these records are reviewed, 

the information enables nursery managers to determine 

if intended improvements actually had a positive effect 

on plant health and growth compared with what was normally done (figure 4.11). More information on trials, or 

experiments is provided in Chapter 20, Discovering Ways 

to Improve Nursery Practices and Plant Quality.

References

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Landis, T.D.; Tinus, R.W.; Barnett, J.P. 1998. The container tree 

nursery manual: volume 6, seedling propagation. Agriculture 

Handbook 674. Washington, DC: U.S. Department of Agriculture, Forest Service. 166 p.

Landis, T.D.; Dumroese, R.K. 2000. Propagation protocols on the 

Native Plant Network. Native Plants Journal. 1: 112–114.

82 Tropical Nursery Manual

Table 4.6—Continued

Hardening

Plant size at start of phase 

(height and stem diameter) 

Average about 10- to 12-cm height, 2.5- to 3.0-mm stem diameter on Nov. 3 

Irrigation (type and frequency; 

for example, daily, every other day, etc.)

Approximately every other day hand-watering or as needed (very hot days will water daily)

Fertilization type, rate, and frequency, if any N/A*

Environmental conditions for crop 

(light, temperature, and humidity)

Full sun, ambient temperature 

Horticultural treatments 

(for example, spacing, cultivation practices)

Some weeding. Weather has been windy; no need to brush. 

Plant size at end of phase 

(height and stem diameter) 

15-cm tall; 3.5-mm stem diameter 

Date hardening phase completed Dec. 10—ready to go

Date plants delivered Dec. 15 

Notes 

(vigor and health, challenges or problems)

Other notes

Notes on performance of crop after outplanting Follow up by phone call scheduled for Jan. 5. Client is using a mattock and shovel to plant.

Figure 4.11—Proper planning ensures that the needs of the 

nursery, the plants, and the clients are met in a timely fashion. 

Photo by Douglass F. Jacobs.

*N/A: Not applicable
Landis, T.D.; Dumroese, R.K. 2002. The Native Plant Network: an 

on-line source of propagation information. International Plant 

Propagators’ Society, Combined Proceedings. 51: 261–264.

Landis, T. D. 2011. The Target Plant concept - a history and brief 

overview. In: Riley, L.E., Haase, D.L. and Pinto, J.R.tech coords. 

National Proceedings: Forest and Conservation Nursery Associations – 2010. Proceedings RMRS-P-65. Fort Collins, CO: US 

Department of Agriculture, Forest Service, Rocky Mountain 

Research Station, Proceedings. 61-66.

Wilkinson, K.M.; Elevitch, C.R. 2003. Growing koa: a Hawaiian legacy 

tree. Holualoa, HI: Permanent Agriculture Resources.

Wilkinson, K.M.; Elevitch, C.R. 2004. Propagation protocol for 

production of container Acacia koa Gray plants: permanent agriculture resources, Holualoa, Hawai‘i. In: Native Plant Network. 

Moscow, ID: University of Idaho, College of Natural Resources, 

Forest Research Nursery. http://www.nativeplantnetwork.org. 

(September 2009).

Wilkinson, K.M.; Elevitch, C.R. 2004. Propagation protocols for 

production of container Cordia subcordata, Thespesia populnea, 

Santalum freycinetianum, Metrosideros polymorpha, and Erythrina sandwicensis species. In: Native Plant N etwork. Moscow, ID: 

University of Idaho, College of Natural Resources, Forest Research 

Nursery. http://www.nativeplantnetwork.org. (September 2009).

Additional Reading

Evans, J. 1992. Forest nurseries. In: Plantation forestry in the 

tropics: tree planting for industrial, social, environmental, and 

agroforestry purposes. Second edition. New York: Oxford University Press. Chapter 10.

Jacobs, D.F.; Wilkinson, K.M. 2008. Planning crops and developing propagation protocols. In: Dumroese, R.K.; Luna, T.; Landis, 

T.D., eds. 2008. Nursery manual for native plants: a guide for 

tribal nurseries. Agriculture Handbook 730. Washington, DC: 

U.S. Department of Agriculture, Forest Service. 302 p

Wescom, R.W. 1999. Nursery production scheduling. In: Wescom, 

R.W. 1999. Nursery Manual for Atoll Environments. RAS/97/330 

Working Paper No. 9. U.S. Department of Agriculture, Natural 

Resources Conservation Service: 12–14. Chapter 5.

Crop Planning: Propagation Protocols, Schedules, and Records 83
84 Tropical Nursery Manual

Appendix 4A—A sample propagation protocol form

Propagation protocol

Family scientific name

Family common name

Scientific name

Common name

General distribution

Propagation goal

Propagation method

Product type

Time to grow

Target specifications

Propagule collection

Propagule processing

Seeds/kg

Purity

Germination

Pre-sowing treatments

Growing area

Growing medium

Establishment phase

Duration of establishment phase

Rapid growth phase

Duration of rapid growth phase

Hardening phase

Duration of hardening phase

Harvesting, storage, and shipping

Outplanting and cultivation comments

Citations
Crop Planning: Propagation Protocols, Schedules, and Records 85

Daily log

Date

Environmental conditions in growing areas 

(light, temperature, humidity, etc.)

Sunrise/Sunset times

Moon phase

Other weather notes 

(cloud cover, etc.)

What water did plants receive? 

(irrigation type and frequency or 

precipitation)

Today’s activities 

(note how many person-hours per activity) 

(fertilization, pest management, transplanting, packing and shipping, making growing 

media, moving crops from one structure to 

another, treating or sowing seeds, and so on)

Growth phase status 

(make notes when a crop moves from one 

phase to another)

Purchases 

(what supplies or equipment were purchased 

and their cost)

Orders 

(what plant materials were 

delivered and payments made)

General crop/nursery observations

Questions or concerns 

Appendix 4B—A sample daily log form
86 Tropical Nursery Manual

Plant development record

Species name

Propagule source

Date(s) of propagule collection

Establishment

Type and length of propagule treatment 

(for example, scarified, stratified)

Date of propagule establishment 

Growing media and tray or container 

type used

Germination notes 

(including date begins and ends, percent germination)

Cutting notes 

(for example, special conditions, hormone treatments)

Date transplanted (if not direct sown) 

Container type and growing media for transplanting

Microorganisms used?

Misting/irrigation 

(type and frequency)

Fertilization type, rate, and frequency, if any 

Environmental conditions for crop 

(light, temperature, humidity)

Horticultural treatments

 (for example, cultivation practices)

Date establishment phase completed

Notes 

(resowing or thinning activities, problems or challenges) 

Appendix 4C—A sample plant development record form
Crop Planning: Propagation Protocols, Schedules, and Records 87

Rapid growth

Time after sowing and sticking to enter rapid growth phase 

Plant size at start of phase (height) 

Container type and growing medium

Irrigation 

(type and frequency; for example, daily, every other day, etc.)

Fertilization type, rate, and frequency

Environmental conditions for crop 

(light, temperature, and humidity)

Horticultural treatments 

(for example, spacing, cultivation practices)

Date rapid growth phase completed

Notes 

(development, vigor and health, challenges or problems)

Hardening

Plant size at start of phase (height and stem diameter) 

Irrigation 

(type and frequency; for example, daily, every other day, etc.)

Fertilization type, rate, and frequency, if any

Environmental conditions for crop 

(light, temperature, and humidity)

Horticultural treatments (for example, 

spacing, cultivation practices)

Plant size at end of phase 

(height and stem diameter) 

Date hardening phase completed

Date plants delivered

Notes 

(vigor and health, challenges or problems)

Other notes

Notes on performance of crop after outplanting

Appendix 4C—Continued
88
89

Many environmental factors influence growth and production of nursery plants. 

The primary processes affected by environmental factors are photosynthesis and transpiration. Photosynthesis is the means by which light energy from the sun is converted 

into chemical energy in the presence of chlorophyll, the green pigment in leaves. During photosynthesis, sugars are produced from carbon dioxide from the air and water 

from the soil while oxygen is released back into the air (figure 5.1). Photosynthesis is 

a “leaky” process because, to allow the intake of carbon dioxide, water vapor is lost 

through pores, or stomata, on the leaf surfaces. This water loss is called transpiration. 

To maximize the photosynthesis necessary for plant growth, growers must reduce the 

factors that limit photosynthesis or increase the factors that promote photosynthesis.

Photosynthesis, and therefore growth, can be limited by factors associated with 

the growing medium, other organisms, or the atmosphere (figure 5.2). Limiting factors related to growing media include water and mineral nutrients and are discussed 

in detail in Chapter 6, Growing Media, Chapter 11, Water Quality and Irrigation, 

and Chapter 12, Plant Nutrition and Fertilization. Photosynthesis can be limited by 

the absence of beneficial organisms or by the presence of harmful organisms; these 

instances are discussed in Chapter 13, Beneficial Microorganisms, and Chapter 14, 

Problem Prevention and Holistic Pest Management. Atmospheric factors that affect 

photosynthesis include light, temperature, relative humidity, and carbon dioxide 

levels. Propagation structures, or environments, are any area that is modified to 

encourage the growth of nursery plants by controlling these atmospheric factors 

during all phases of growth in the nursery. Understanding different types of propagation structures and how they work is critical whether designing a new nursery 

facility or modifying an existing one.

Tropical nursery facilities differ not only in their complexity but also in their 

biological and economical aspects. Propagation environments can be as simple as a 

garden plot where water and fertilizer are applied, or as complex as high-tech greenhouses that also modify all atmospheric factors. Although modifying atmospheric 

factors can also influence the growing media and occurrence of other organisms, our 

focus in this chapter is primarily on modifying atmospheric factors (light, temperature, and relative humidity) through a variety of propagation environments.

Propagation Environments

Douglass F. Jacobs, Thomas D. Landis, Tara Luna, and Diane L. Haase 5

Facing Page: Propagation environments modify atmospheric factors such as light and temperature. In this nursery, removable 

palm thatch provides shade for young seedlings but allows full sun to reach maturing plants. Photo by Thomas D. Landis.
Matching Propagation 

Environments to the Site

The best possible propagation environment is designed 

for a specific nursery site (Landis 1994). Whether building a new nursery or modifying an existing one, it is 

critical to analyze the limiting factors on the site. The 

value of such an analysis can be clearly demonstrated on 

the Big Island of Hawai‘i. One of the Hawai‘i Volcanoes 

National Park Service nurseries is located near sea level 

where the most limiting site factors are intense sunlight 

and seasonal high winds. This situation requires a strong 

structure to withstand wind loads and a shadecloth to 

minimize heat buildup. Moving up slope to the town 

of Volcano at 3,700 ft (1,130 m), the climate is milder 

year round than at the coast and the limiting factors are 

low sunlight with heavy rains, so a structure with a clear 

covering to maximize light transmission while protecting the crop from heavy rains would be ideal. Moving to 

an even higher elevation (7,000 ft [2130 m]), the limiting factor at Hakalau National Wildlife Refuge nursery is 

cold, with nighttime temperatures that can dip to freezing any time of the year. Thus, it is necessary to have an 

enclosed structure to protect the crops. As expected, the 

costs of nursery development increase with greater control of the propagation environment. A nursery that is 

well matched to its environment, however, will be much 

less expensive to operate than a poorly designed one.

The Challenge of Growing Many 

Species and Stocktypes

Tropical nurseries grow an increasing diversity of 

native plant species. Often, these diverse crops must be 

started on various dates, so, at any one time, a nursery 

might have everything from germinating seeds to large 

plants. Although some species are grown from seeds, 

others in the same nursery might have to be grown from 

rooted cuttings. So, a good native plant nursery should 

be designed with various propagation environments in 

which plants of similar requirements and growth stages 

can be grown. For example, a controlled environment 

such as a greenhouse or other sheltered area can provide 

ideal conditions for germinating seeds and establishing 

young seedlings in containers. After young seedlings are 

established, they could be moved to a shadehouse or open 

compound to continue their growth. During hardening, 

the crops can be acclimated to the ambient environment 

in the same shadehouse or open compound.

90 Tropical Nursery Manual

Figure 5.1—Two important processes occur in the leaves of 

green plants. In photosynthesis, sunlight triggers a chemical reaction in which water from the soil and carbon dioxide from the 

air are converted to sugars and oxygen, which are released back 

to the atmosphere. During the process, water vapor is lost from 

the leaves in a process known as transpiration. Illustration from 

Dumroese and others (2008).

Figure 5.2—It is useful to think of the nursery environment in 

terms of factors that might be limiting to plant growth. Limiting 

factors in the soil include water and mineral nutrients whereas, 

temperature, light, carbon dioxide, and humidity can be limiting 

factors in the atmosphere. Other organisms can either be beneficial 

or detrimental. Illustration from Dumroese and others 2008.
Minimally Controlled Propagation 

Environments

A minimally controlled environment is the simplest and 

least expensive of all types of propagation environments. 

The most common type is an open growing compound. It 

consists of an area where plants are exposed to full sunlight 

and is usually nothing more than an irrigation system and 

a surrounding fence.

Open Growing Compounds

Open growing compounds are popular in tropical climates (figure 5.3). Nurseries use open compounds for plant 

propagation and for areas to expose crops previously grown 

inside structures to ambient conditions during hardening. 

Plants can be grown on elevated platforms, benches, or pallets to improve air pruning of the roots, or directly on a layer 

of gravel (to provide drainage) that is covered with landscape 

fabric (to control weeds). Irrigation is provided by sprinklers 

for smaller containers or driplines for larger ones; plants 

obtain nutrients from controlled-release fertilizers that are 

incorporated into the growing media. The compound needs 

to be fenced to minimize animal damage, and, in windy areas, 

a shelterbelt of trees around the compound can protect from 

desiccation and improve irrigation coverage.

Wetland Ponds

Artificial ponds are another type of minimally controlled 

environment. They are used for growing riparian and wetland plants and are especially good for propagating sedges 

and rushes. They can also be used to provide specific habitats 

for certain wetland plants, such as for mangroves adapted to 

saline coastal habitats. Wetland ponds can be aboveground 

reservoirs, such as children’s wading pools or cattle troughs, 

or they can be constructed with pond liners either in an 

excavated area or at ground level using a raised perimeter 

(figure 5.4). These simple propagation environments use 

growing media amended with controlled-release fertilizer 

and require only periodic flood irrigation. Some islands, like 

Hawai‘i and American Samoa, have freshwater wetland species that can be grown as container plants with freshwater 

flood irrigation.

Propagation Environments 91

Figure 5.3—Open compounds, like Waimea State Tree Nursery on the Big Island of Hawai‘i, are common in tropical climates and often 

used for hardening crops grown in greenhouses or other structures (A). Open compounds often have irrigation and are fenced as seen at 

this Republic of Palau Forestry Department nursery (B). Photo A by Thomas D. Landis, and photo B by Tara Luna.

Figure 5.4—Wetland ponds can be constructed in the outdoor 

nursery for growing wetland species. On a smaller scale, plastic 

tubs can be used instead. Photo by Thomas D. Landis.

A B
Semicontrolled Propagation 

Environments

Another category of propagation environments is called 

“semicontrolled” because only a few of the limiting factors 

in the ambient environment are modified. Semicontrolled 

environments consist of a wide variety of growing structures ranging from simple cold frames to shadehouses.

Cold Frames

Cold frames are low-to-the-ground structures consisting of a wood or metal frame with a transparent covering. 

As their name suggests, they have no heating source except 

for the sun. Cold frames are the most inexpensive propagation structure and are easy to build and maintain. Because 

conditions inside can stay relatively warm and moist, cold 

frames can be used for seed germination or rooting cuttings. They can also be used to protect seedlings and cuttings from heavy rains and wind.

The ideal location for a cold frame is an area with a 

slight slope to ensure good drainage. A sheltered spot 

against the wall of a building or greenhouse provides 

additional protection (figures 5.5A). Some nurseries sink 

the floor of the cold frame 6 to 12 in (15 to 30 cm) into the 

ground to use the earth for insulation. Other nurseries 

make their cold frames lightweight enough to be portable 

so they can move them from one section of the nursery 

to another.

It is relatively easy to build a cold frame. Frames are 

usually made of wood such as mahogany that will resist 

decay; the new recycled plastic lumber also works well. 

Never use creosote-treated wood or wood treated with 

pentachlorophenol because these substances are toxic 

to plants. The cold frame needs to be built so that it is 

weather-tight and so the top can be opened partially or 

fully to allow for various levels of ventilation, watering, 

and the easy removal of plants (figure 5.5B, 5.5C). The 

cover must be able, however, to be attached securely to 

the frame to resist wind gusts. Heavy plastic film is an 

inexpensive covering but usually lasts only a single season. Hard plastic or polycarbonate panels are more durable and will last for several years. Cold frame kits may 

also be purchased and are easily assembled; some kits 

even contain automatic ventilation equipment.

Cold frames can be labor-intensive because they need 

to be opened and closed daily to manage temperature and 

humidity levels. A thermometer that can be conveniently 

read without opening the cover is mandatory. In a cold 

frame, plants grow best at 65 to 85 °F (18 to 29 °C). If air 

temperature goes above 85 °F (29 °C), the top must be 

opened to allow ventilation. In the tropics, cold frames usually need to have shadecloth suspended above them to help 

moderate temperatures.

Hoop Houses and 

Polyethylene Tunnels

Hoop houses and polyethylene (“poly”) tunnels are 

versatile, inexpensive propagation environments. They 

are usually constructed of semicircular frames of polyvinyl chloride (PVC) or metal pipe covered with a single 

layer of heavy polyethylene and are typically quite long 

(figure 5.6). Some hoop houses have end walls made 

of solid material such as water-resistant plywood. The 

cover on hoop houses is changed or removed during the 

growing season to provide a different growing environment, 

eliminating the need to move the crop from one structure to 

another. In general, during the rainy season, a clear plastic 

92 Tropical Nursery Manual

Figure 5.5—Cold frames are an inexpensive alternative to a 

greenhouse. Cold frames should be placed in a sheltered location for additional protection (A). Coverings may be removed 

(B) or held open to manage humidity and heat levels (C). Photos 

by Tara Luna.

B

C

A
cover is used during seed germination and seedling establishment. The plastic cover protects the young germinants 

and seedlings from heavy rains and can be pulled back as 

needed to provide adequate ventilation. As the dry season 

approaches and the seedlings are established, the plastic 

cover can be removed and replaced with shadecloth. Sometimes, a series of shadecloths, each with a lesser amount of 

shade, are used to gradually expose crops to full sun. During hardening, the shadecloth is completely removed to 

expose the plants to the ambient environment.

Shadehouses

Shadehouses are the most permanent of semicontrolled 

propagation environments and serve several uses. In the 

tropics, shadehouses are commonly used to propagate 

Propagation Environments 93

Figure 5.6—Hoop houses can be used for a variety of propagation environments by changing or removing the coverings. Photo 

by Douglass F. Jacobs.

Figure 5.7—Shadehouses are semicontrolled environments that are used for protecting plants from intense sunlight, rain, or wind (A). 

They can be constructed of wood frames with lath or metal frames with shadecloth and are equipped with irrigation systems that can also 

apply liquid fertilizer (B). In addition, shade can be created with locally available materials (C). Photos by Tara Luna.

plants under conditions of intense sunlight (figure 5.7A). 

Less permanent shadehouses consist of a metal pipe frame 

covered with shadecloth and allow for rapid take down 

when tropical storms threaten. When used for growing, 

shadehouses can be equipped with sprinkler irrigation 

and fertilizer injectors (figure 5.7B). When the shade is 

installed on the sides of the structure, shadehouses are very 

effective at protecting crops from wind and therefore help 

to reduce transpiration. Shadehouses can also be built with 

local materials (figure 5.7C).

Fully Controlled Propagation 

Environments

Fully controlled environments are propagation structures in which all or most of the limiting environmental 

factors are controlled. Examples include growth chambers 

(high-cost option used almost exclusively for research) 

and greenhouses. Tropical nurseries with large forestry 

and restoration programs often make use of greenhouses. 

Fully controlled environments have the advantage of yearround production in nearly any climate. In addition, most 

crops can be grown faster and have more uniform quality 

than those grown in propagation environments with less 

control. These benefits must be weighed against the higher 

costs of construction and operation. The more complicated 

a structure is, the more problems that can develop. This 

concept is particularly true in the remote locations of many 

tropical nurseries, where electrical power outages can be 

common and it is difficult, time-consuming, and expensive to obtain specialized parts and repair services. Some 

good references that provide more detail about greenhouse 

design include Aldrich and Bartok (1989), Bartok (2000), 

and Landis and others (1994).

A B C
Greenhouse Engineering 

Considerations

All greenhouses are transparent structures that allow 

natural sunlight to be converted into heat (figure 5.8A). 

At the same time, greenhouses are poorly insulated and 

require specialized cooling and ventilation systems to regulate temperatures. Keeping a greenhouse cool during sunny 

days requires carefully engineered ventilation systems. It 

is important to understand that greenhouse construction 

requires specialized skills (figures 5.8B, 5.8C). In developed areas, greenhouses may be regulated by municipal 

building codes and zoning, which is another good reason 

to work with a professional contractor before buying and 

constructing a greenhouse.

Design Loads

The load on a greenhouse includes dead loads (the 

weight of the structure), live loads (caused by building use), 

and weather-related loads (wind). Live loads include people working on the structure and the weight of equipment, 

such as irrigation systems, fans, lighting systems, and even 

hanging plants.

Foundations, Floors, and Drainage

The foundation connects the greenhouse to the 

ground and counteracts the design load forces. Inexpensive floors can consist of gravel covered with landscape 

cloth, but the ground beneath the floor must drain water 

freely to prevent standing water and safety issues. Nursery crops require frequent irrigation and, depending on 

the irrigation system, much of this water may end up on 

the floor. Drain tiles might be needed to ensure that the 

greenhouse floor will not become a swamp. Full floors can 

be engineered with drains so that all water and fertilizer 

runoff can be contained on site; runoff containment is a 

legal requirement in many places. It may be necessary to 

design the greenhouse so that all wastewater drains into 

a pond or constructed wetland to prevent contamination 

of water sources; these ponds or wetlands can sometimes 

be used for other purposes, such as growing wetland 

plants as described previously in this chapter.

If wheeled equipment will be used to move plants, 

concrete walkways between the benches are necessary. 

Note that black asphalt heats up rapidly and becomes 

soft, so concrete is a better, but more expensive, option. 

Full concrete floors will eliminate many pest problems, 

especially algae, moss, and liverworts that thrive in the 

humid nursery environment.

94 Tropical Nursery Manual

Figure 5.8—Greenhouses, like this one in American Samoa, are 

the most sophisticated propagation environments (A). Workers 

with specialized skills are needed from the initial surveying (B) 

to the final construction (C). Photo A by Thomas D. Landis, and 

photos B and C by Ronald Overton.

B

C

A
Framing Materials

Ideal framing supports the greenhouse covering with 

minimal shading and heat loss while allowing ease of access 

and handling. Framing materials include galvanized steel, 

aluminum (lightweight but expensive), and treated wood.

Greenhouse Kits

The heating and cooling systems of fully controlled greenhouses must be carefully engineered to match both the size 

of the structure and the ambient environment. Be aware that 

inexpensive greenhouse kits often have vents or fans that are 

too small for the size of the greenhouse. Kit greenhouses were 

designed for some “average” environment and will probably 

have to be modified to handle the limiting environmental 

factors on your site. Before purchasing a greenhouse kit, it is 

a good idea to hire an experienced consultant, speak with a 

knowledgeable company representative, and discuss designs 

with other growers or professionals.

Greenhouse Coverings 

A wide variety of greenhouse coverings are available 

and the selection of a particular type is usually based on 

cost, type of structure, and the environmental conditions at 

the nursery site (figure 5.9).

Polyethylene tarps are relatively cheap but require 

replacement every 2 to 4 years depending on the grade 

of plastic. Double layers of polyethylene sheeting that are 

inflated with a fan are stronger and provide better insulation 

longer than a single layer. The two layers are attached to the 

framing with wooden furring strips or specially designed 

fasteners. This process is relatively simple so many growers 

change their own coverings. Because they are so well insulated and airtight, polyethylene greenhouses require good 

ventilation to prevent condensation.

Polycarbonate (“polycarb”) panels, the most popular 

permanent greenhouse covering, have about 90 percent of 

the light transmission properties of glass. Polycarb is strong 

and durable but is one of the more expensive coverings.

These panels are the most common greenhouse coverings, and a more detailed description of costs and engineering and operational considerations is available in Landis 

and others (1994).

Controlling Greenhouse Temperatures

One of the most challenging aspects of a greenhouse is 

controlling temperature. Sunlight is converted into heat that 

can become lethal to plants. A sophisticated control system 

that can maintain a designated temperature through a series 

of heating and cooling stages is a necessary and wise investment to minimize energy costs. Vents and fans are used to 

keep air moving inside the greenhouse and exhaust heat 

from the structure. In dry, windy environments, wet walls 

use the power of evaporation to cool incoming air. Growers 

can also use short bursts of their irrigation system for cooling. 

Automatic sensing instruments are available that can be connected with cooling equipment to trigger a cooling cycle for 

the greenhouse. Mechanical thermostats provide the best and 

most economical form of temperature control and can be used 

to activate motorized vents and fans within a greenhouse.

Specialized Rooting Propagation 

Environments

The most common type of vegetative propagation is the 

rooting of cuttings. Often, this form of propagation requires 

a specialized environment known as a rooting chamber that 

creates specific conditions to stimulate root initiation and 

development. Because cuttings do not have a root system 

(figure 5.10A), rooting chambers must provide frequent 

misting to maintain high humidity to minimize transpiration. Root formation is stimulated by warm temperatures 

and moderate light levels; these conditions maintain a high 

level of photosynthesis. Therefore, many rooting chambers 

are enclosed with polyethylene coverings that, in addition 

to maintaining high humidity, keep the area warm. If the 

chambers are outside, the covering further protects cuttings 

from rain and drying winds. For more information on rooting cuttings, see Chapter 10, Vegetative Propagation.

Enclosed Rooting Chambers

Because it is easy to construct and very economical, a 

simple enclosed rooting chamber is essentially the same 

Propagation Environments 95

Figure 5.9—Greenhouses are covered with transparent coverings such as plastic sheeting or hard plastic panels to maximize 

the amount of sunlight reaching the crop. Photo by Brian F. Daley.
as the cold frame discussed earlier. They are commonly 

used in tropical nurseries throughout the world because 

they do not require electricity. Enclosed rooting chambers 

rely on manual operation so they require diligent daily 

inspection to regulate humidity and air temperatures, and 

the rooting medium must be watered as needed. They typically have shadecloth suspended above them to moderate 

temperatures, but, if heat or humidity becomes excessive, 

enclosed chambers need to be opened for ventilation. One 

design is known as a “poly propagator” because it is covered with polyethylene plastic sheeting and is a simple and 

inexpensive design (figure 5.10B).

Intermittent-Mist Rooting Systems

These rooting propagation environments are either 

enclosed or open. Rooting cuttings is much easier in these 

environments because intermittent-mist rooting chambers have a high degree of environmental control (figure 

5.11A). Clock timers (figure 5.11B) control the timing 

and duration of misting from specialized nozzles (figure 

5.11C). Frequent misting maintains high humidity that 

reduces water loss from the cuttings, and evaporation of 

the small droplets moderates air and leaf temperature.

Mist systems require high water pressure that is supplied through PVC pipes. Mist timing is controlled by two 

time clocks that open and close a magnetic solenoid valve 

in the water line linked to the nozzles. One clock turns the 

system on during the day and off at night and the other 

controls the timing and duration of the mists. Because the 

aperture of the mist nozzles is so small, a cartridge filter 

needs to be installed in the water line after a gate valve.

Because of the proximity of water and electricity, all 

employees need to receive safety training. All wiring used 

for mist propagation must be grounded and must adhere 

to local building codes. Electrical outlets and components must be enclosed in waterproof coverings. The high 

humidity encourages the growth of algae and mosses, so 

the mist propagation system needs to be cleaned regularly. Mist systems require water that is low in dissolved 

salts; “hard” water may result in whitish deposits that 

can plug mist nozzles. See Chapter 11, Water Quality and 

Irrigation, for more information.

96 Tropical Nursery Manual

Figure 5.10—Rooting cuttings require a specialized propagation environment because cuttings lack a root system (A). The “poly 

propagator” is the most simple and inexpensive rooting chamber (B). Illustrations by R.H.F. Wilson from Longman 1993.

A B
Modifying Light and Temperature 

in Propagation Environments

As mentioned earlier, light is necessary for photosynthesis, which provides energy for plant growth. For lightloving species, more light results in more growth (figure 

5.12A), but greenhouse light levels are often too intense to 

grow some species of tropical plants (table 5.1). As a result, 

growers apply shadecloths to lessen light intensity and 

the resultant heat (figures 5.12B, 5.12C). Shadecloths are 

rated by the amount of shade they produce, ranging from 

30 to 80 percent. Black has been the traditional color 

because it is relatively inexpensive, but now shadecloth 

comes in white, green, and reflective metal. Because black 

absorbs sunlight and converts it into heat that can be conducted into the propagation structure (figure 5.12D), black 

shadecloth should never be installed directly on the covering of any propagation structure, but instead needs to be 

suspended above it to facilitate air movement. Although 

more expensive than black shadecloth, white or aluminized 

shade fabrics are better for tropical environments and will 

do a much better job of cooling the propagation environment while still keeping light levels high. Applying a series 

of shadecloths, each with a lesser amount of shade, over a 

period of time is a good way to gradually harden nursery 

stock and prepare it for outside conditions.

Monitoring Temperatures

Thermometers that record the maximum and minimum 

temperatures during the day are simple and economical 

instruments (figure 5.13A) that can help growers monitor subtle microclimates within any propagation environment. New devices, such as self-contained, programmable 

temperature sensors, are revolutionizing the ways in which 

temperature can be monitored in nurseries (figures 5.13B, 

5.13C). Many of these sensors are small enough to be placed 

within a container or storage box and can record temperatures (between -40 and 185 °F [-40 and 85 °C]) for more 

than 10 years. Because these single-chip recording devices 

can be submersed in water and are resistant to dirt and 

impact, they can be used to monitor temperatures under 

Propagation Environments 97

Table 5.1—Shade requirements of a variety of tropical 

plant species.

Figure 5.11—Intermittent-mist systems are easily controlled environments (A). Programmable timers control the timing and duration 

(B) of specialized mist nozzles (C), which keep humidity levels high, reduce transpiration, and provide cooling. Photos A and C by Tara 

Luna, and photo B by Thomas D. Landis.

Scientific name Light 

requirement 

Acacia species Sun

Pandanus species Sun

Scaevola species Sun

Bobea species Partial shade

Bonamia menziesii Partial shade

Pritchardia species Partial shade

Cyathea, Cibotium species Shade

Cyrtandra species Shade

Elaeocarpus bifidus Shade

A B C
98 Tropical Nursery Manual

Figure 5.12—Sunlight provides the energy necessary for plant growth (A) but is converted to heat inside propagation structures. 

Shadecloth reduces light intensity and cools the environment in small enclosures (B) and larger shadehouses (C). Compared with 

white or reflective shadecloth, black shadecloth can absorb heat and radiate it into the propagation environment (D). Illustration A 

from Dumroese and others (2008), photo B by Tara Luna, photo C by Brian F. Daley, and illustration D by Jim Marin.

Figure 5.13—Monitoring and controlling temperature is critical to successfully growing a crop of 

tropical plants (A). Many nurseries use small, programmable, self-contained temperature sensors 

that record daily maximum and minimum temperatures (B and C). Photo A by Thomas D. Landis, 

and photos B and C by David E. Steinfeld.

B

A C

D

A B C
most nursery conditions. The data recorded on the sensors 

must be downloaded to a computer and can then be easily 

placed into a spreadsheet. The small size of the sensor can 

also be a drawback; it is easy to misplace. Attach a strip of 

colorful flagging to indicate where the sensors are located 

and write any necessary information on the flagging with a 

permanent marker.

Equipment Maintenance

Even if you purchase the most reliable “automatic” 

environmental control equipment, it must be monitored 

and maintained. The hot and humid tropical nursery environment is particularly hard on equipment, and especially 

so in coastal areas where equipment may also be exposed to 

salt. Regular maintenance ensures longevity, reduces costly 

repairs, and may help avoid disasters.

Routine maintenance of all greenhouse and nursery 

operation equipment should be a top priority. Someone 

who is mechanically inclined should be given the responsibility for equipment maintenance. Equipment records 

should be included in the nursery’s daily logbook. See 

Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records, and Chapter 19, Nursery Management, 

for more details. These log books will be invaluable when 

solving problems, budgeting, and developing maintenance 

plans. A system of “promise cards” specifies when servicing 

needs to be done and can be incorporated into the nursery 

computer system. Keep a supply of spare parts on hand, 

especially parts that may not be readily available or may 

take a long time to receive. It is a good idea to have a spare 

cooling fan motor on standby and a handy supply of hardware items such as washers, screws, and bolts. Familiarize 

all employees on the operation of all equipment so that 

problems can be detected early. The instruction manuals 

for all equipment need to be kept and easy to find.

References

Aldrich, R.A.; Bartok, J.W., Jr. 1989. Greenhouse engineering. 

NRAES-33. Ithaca, NY: Cornell University, Northeast Regional 

Agricultural Engineering Service. 203 p.

Bartok, J.W., Jr. 2000. Greenhouse for homeowners and gardeners. 

NRAES-137. Ithaca, NY: Cornell University, Northeast Regional 

Agricultural Engineering Service. 200 p.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Landis, T.D. 1994. Using “limiting factors” to design and manage propagation environments. International Plant Propagators’ 

Society, Combined Proceedings. 43: 213–218.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1994. 

The container tree nursery manual: volume 1, nursery planning, 

development, and management. Agriculture Handbook 674. 

Washington, DC: U.S. Department of Agriculture, Forest Service. 

188 p.

Longman, K.A. 1993. Tropical trees: propagation and planting 

manuals: volume 1, rooting cuttings of tropical trees. London, 

United Kingdom: Commonwealth Science Council. 137 p.

Additional Reading

Clements, S.E.; Dominy, S.W.J. 1990. Costs of growing containerized seedlings using different schedules at Kingsclear, New 

Brunswick. Northern Journal of Applied Forestry. 7: 73–76.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1992. The 

container tree nursery manual: volume 3, atmospheric environment. Agriculture Handbook 674. Washington, DC: U.S. Department of Agriculture, Forest Service. 145 p.

Propagation Environments 99
100
101

A growing medium can be defined as a substance through which plant roots 

grow and extract water and nutrients. Selecting a good growing medium is fundamental to good nursery management and is the foundation of a healthy root 

system.

Growing media for use in container nurseries is available in two basic forms: 

soil based and organic based. Compared with soil based media that has field soil 

as a major component, organic based media (a base of organic materials that 

may be compost, peat, coconut coir, or other organic materials, mixed with 

inorganic ingredients) promotes better root development. In temperate areas, 

nurseries can choose from a wide range of commercial products for their growing media, including peat moss, vermiculite, and perlite, and premixed blends 

of these ingredients. Most nurseries in the tropics, however, do not have easy 

and affordable access to these materials, and even nurseries in temperate areas 

are seeking to replace some of these ingredients with more local and sustainable materials. In the tropics, growers often create their own media using locally 

available ingredients.

A favorable growing medium consists of two or more ingredients. Growers must be familiar with the positive and negative characteristics of the various ingredients and how they will affect plant growth when creating a suitable 

growing medium, or even when purchasing a commercial one. This chapter 

describes the uses, functions, and properties of growing media ingredients. From 

this information, you can experiment with available materials and find the best 

combination(s) for your nursery.

Growing Media

Thomas D. Landis, Douglass F. Jacobs, Kim M. Wilkinson, and Tara Luna 6

Facing Page: Many tropical nurseries create their own growing media with a mix of local materials. Photo by Douglass F. Jacobs.
Functions of Growing Media

A growing medium serves four functions (figure 6.1).

1. Physical Support

The growing medium must be porous yet provide 

physical support. Young plants are fragile and must 

remain upright so that they can photosynthesize and 

grow. With larger nursery stock in individual containers, a growing medium must be heavy enough to hold 

the plant upright against the wind. Bulk density is the 

responsible factor and will be discussed in the next section.

2. Aeration

Plant roots need a steady supply of oxygen to convert 

the photosynthate from the leaves into energy so that the 

roots can grow and take up water and mineral nutrients. 

The byproduct of this respiration is carbon dioxide that 

must be dispersed into the atmosphere to prevent the 

buildup of toxic concentrations within the root zone. This 

gas exchange occurs in the large pores (macropores) or 

air spaces in the growing medium. Because nursery plants 

grow rapidly, they need a medium with good porosity—a 

characteristic termed “aeration” that will be discussed in 

more detail in the next section.

3. Water Supply

Nursery plants use a tremendous amount of water for 

growth and development, and this water supply must be 

provided by the growing medium. Growing media are 

formulated so that they can hold water in the small pores 

(micropores) between their particles. Many growing media 

contain a high percentage of organic matter such as peat 

moss and compost because these materials have internal 

spaces that can hold water like a sponge. Therefore, growing media must have adequate porosity to absorb and store 

the large amounts of water needed by the growing plant.

4. Supply of Mineral Nutrients

Most of the essential mineral nutrients that nursery 

plants need for rapid growth must be obtained through the 

roots from the growing medium. Most mineral nutrients 

are electrically charged ions. Positively charged ions (cations) include ammonium nitrogen (NH4

+), potassium (K+), 

calcium (Ca+2), and magnesium (Mg+2). These cations are 

attracted to negatively charged sites on growing medium 

particles up to the point when the roots extract the cations. 

The capacity of a growing medium to adsorb these cations 

is referred to as cation exchange capacity (CEC), and this 

important characteristic is discussed in the next section. Different media components vary considerably in their CEC, 

but peat moss, vermiculite, and compost have a high CEC 

value, which explains their popularity in growing media.

102 Tropical Nursery Manual

Figure 6.1—Primary functions of 

growing media include the capacity 

to hold water and nutrients for root 

uptake, providing adequate root 

aeration, and ensuring structural 

support to the plant. Illustration 

from Dumroese and others (2008).
Physical Properties of 

Growing Media

Water-Holding Capacity

Micropores absorb water and hold it against the pull of 

gravity until plants can use it (figure 6.2). The water-holding capacity of a medium is defined as the percentage of 

total pore space that remains filled with water after gravity drainage. A good growing medium has a high waterholding capacity but also contains enough macropores to 

allow excess water to drain away and prevent waterlogging. Water-holding capacity varies by the types and sizes 

of the growing medium ingredients. For example, a peat 

moss particle will hold much more water than a similarly 

sized piece of pumice. The degree of compaction is also 

extremely important. When growing medium particles are 

damaged during mixing or compacted when the containers 

are filled, the percentage of macropores is severely reduced. 

Overmixed or compacted media will hold too much water 

and roots will suffocate. Finally, the height of the container 

affects the water-holding capacity; a certain amount of 

water will always remain in the bottom of the container 

(figure 6.2). When filled with the same medium, short containers will have a higher percentage of waterlogging than 

taller ones (see Chapter 7, Containers).

Aeration

The percentage of pore space that remains filled with 

air after excess water has drained away is known as aeration. As we have already discussed, oxygen for good healthy 

roots is supplied through the larger macropores (figure 

6.2), which also allow the carbon dioxide from respiration 

to dissipate. A good growing medium, especially for rooting cuttings, contains a high percentage of macropores.

Porosity

The total porosity of a growing medium is the sum of 

the space in the macropores and micropores; plants need 

both. A growing medium composed primarily of large particles will have more aeration and less water-holding capacity than a medium of smaller particles, which will have less 

aeration and more water-holding capacity (figure 6.3). 

Either of these media would restrict plant growth. Plants 

growing in a medium with all large particles would dry out 

too quickly, and those growing in a medium with all small 

particles would suffer from waterlogging. For a singlecomponent medium, the ideal particle range to promote 

both water-holding capacity and aeration is about 0.03 to 

0.24 in (0.8 to 6 mm). In actual practice, however, a good 

growing medium will contain a mixture of ingredients 

with different particle sizes and characteristics.

Bulk Density

Media bulk density is the weight per volume and varies 

with the inherent bulk density of its ingredients and how 

much they are compressed. An ideal growing medium is 

lightweight enough to facilitate handling and shipping 

while still having enough weight to provide physical support. For a given container type and growing medium, 

excessive bulk density indicates compaction. Bulk density and porosity are inversely related; when bulk density 

increases, porosity decreases. Even a very porous growing 

medium can be ruined if it is compressed when the containers are filled.

Chemical Properties of 

Growing Media

Fertility

Rapidly growing young plants use up the stored nutrients in their seeds soon after emergence. Thereafter, plants 

must rely on the growing medium to meet their increasing 

demands for mineral nutrients. As described in Chapter 12, 

Plant Nutrition and Fertilization, many container nursery 

managers prefer media with inherently low fertility (for 

Growing Media 103

Figure 6.2—A good growing medium contains micropores 

that hold water and macropores that allow for air exchange. All 

containers also have a perched water table in the bottom (see 

also Figure 7.2). Adapted from Landis and others (1989).
example, peat-vermiculite) to discourage damping-off during the establishment phase and add soluble fertilizers to 

media throughout the remainder of the growing season. If 

fertilizers are difficult to obtain or cost prohibitive, organic 

amendments such as manure or compost can be included 

in the growing medium. Some plants grow better under 

low fertilization; in addition, beneficial microorganisms, 

such as mycorrhizal fungi, sometimes require low fertility 

to become established on plant roots. See Chapter 13, Beneficial Microorganisms, for more discussion on this topic.

pH

The pH of growing medium is a measure of its relative 

acidity or alkalinity. pH values range from 0 to 14; those 

below 7 are acidic and those above 7 are alkaline. Most 

native plants tend to grow best at pH levels between 5.5 and 

6.5, although some species are tolerant of higher or lower 

pH levels. The main effect of pH on plant growth is its control on nutrient availability (figure 6.4). For example, phosphorus availability drops at extreme pH values because 

phosphorus binds with iron and aluminum at low pH levels and with calcium at high pH levels. The availability of 

micronutrients, such as iron, is even more affected by pH. 

Iron chlorosis, caused by high pH, is one of the most common nutrient deficiencies of nursery stock. Exceptionally 

104 Tropical Nursery Manual

Figure 6.4—The availability of all mineral nutrients is affected 

by the pH of the growing medium. In organic-based growing media, 

maximum availability occurs between 5.5 and 5.6. Illustration 

adapted from Bunt (1988) by Jim Marin.

Figure 6.3—These two containers each contain growing media 

with either large (left) or small (right) particle sizes (A). The corresponding balloons show the relative amounts of air being held, and 

the glasses show the relative amounts of water being held. When 

particle sizes are too small, such as this example of a high-clay growing medium, then root development is inhibited (B). Illustration A 

by Jim Marin, and photo B by J.B. Friday.

B

A
high or low pH levels also affect the abundance of pathogens and beneficial microorganisms. For example, low pH 

can predispose young plants to damping-off fungi.

Cation Exchange Capacity (CEC)

CEC refers to the ability of a growing medium to hold 

positively charged ions. Because most growing media are 

inherently infertile, CEC is a very important consideration. 

In the growing medium, plant roots exchange excess charged 

ions for charged nutrient ions (figure 6.1), and then these 

nutrients are transported to the foliage, where they are used 

for growth and development. Because the CEC of a growing 

medium reflects its nutrient storage capacity, it provides an 

indication of how often fertilization will be required. Because 

nutrient leaching occurs during irrigation, container nurseries prefer a growing medium with a very high CEC.

Biological Properties of 

Growing Media

Growing media may contain pathogenic bacteria or 

fungi. Growing media ingredients that may contain pathogens can be treated with sterilization or pasteurization before 

use, as described later in this chapter. Organic-based growing 

media are preferred in nurseries because they are generally 

pest free. Although peat moss is not technically sterile, it 

does not contain pathogens or weed seeds when obtained 

from reliable sources. Vermiculite and perlite are rendered 

completely sterile during manufacturing, when they are 

exposed to temperatures as high as 1,832 °F (1,000 °C). 

Well-prepared composts are generally pest free because 

sustained, elevated temperatures during composting kill 

most pathogens. Another benefit of composting is that 

beneficial microorganisms increase in the final stages of the 

process. Composted bark of some tree species, for example, 

contains microbes that suppress common fungal pathogens and nematodes. These suppressive effects depend on 

the parent material and composting time (Castillo 2004).

Growing Media Ingredients

Once the functions and characteristics of growing media 

ingredients are understood (table 6.1), an effective and 

affordable growing media can be developed. Many tropical 

nurseries mix growing media themselves. Some purchase 

premixed commercial brands. Some may mix their own 

media for larger containers, but purchase sterile media for 

germinants. A typical growing medium is a composite of 

Growing Media 105

Table 6.1—Different chemical and physical properties of some common materials used to create growing media. Soils are covered separately because they are a combination of both organic and inorganic components, and are extremely variable. Adapted from Buamscha and 

Altland (2005), Johnson (1968), Lovelace and Kuczmarski (1994), and Newman (2007).

Component Bulk density Porosity:water Porosity: air pH Cation exchange capacity

Organic ingredients

Sphagnum peat moss Very low Very high High 3 to 4 Very high

Bark Low Low Very high 3 to 6 High

Coir Low High High 6 to 7 Low

Sawdust Low High Moderate 3 to 6 Low

Rice hulls Low Low Moderate 5 to 6 Low

Compost Variable Variable Variable 6 to 8 High

Inorganic ingredients

Vermiculite Very low Very high High 6 to 8 High

Perlite Very low High High 6 to 8 Very low

Sand Very high Moderate Very low Variable Low

Pumice Low Low High 6 to 8 Low

Field soil

Field soil Variable Variable Variable Variable Variable
Why Field Soil Is a Poor Growing Medium 

Although it may seem intuitive that potting up nursery plants in containers filled with local field soil would work 

well, it is not the case. Placing soil in a container results in growing conditions that are completely different from those 

of unrestricted field soil. These conditions are unfavorable to healthy root growth and plant development in the nursery. In fact, field surveys of tropical and sub-tropical areas of Africa, Asia, and Latin America showed unexpectedly 

poor field growth of seedlings after outplanting, and correlated this problem with poor root development because of 

using soil-based media in nurseries (Miller and Jones 1995). The recommended alternative is to replace soil-based 

media whenever possible with an organic-based, or “artificial” growing media consisting of ingredients such as compost, peat, or coconut coir (Miller and Jones 1995). If soil must be used, it should be only a small percentage of the mix, 

amended with other ingredients to overcome some of the problems listed in the following sections.

Restricted Volume

Unlike in field conditions, nursery plants will have access to a very limited amount of soil. This limited rooting volume provides seedlings with only small reserves of available water and nutrients and the amount of these resources can change quickly. 

Because plants will only have a limited area in which to grow their roots, this medium needs to be the best possible material.

Imbalance of Soil Microorganisms

Native soils contain a myriad of microorganisms: some are beneficial and some are pathogenic. These organisms exist in a 

natural balance in field soils, but when these soils are placed in the nursery growing environments, this delicate balance is upset 

and problems can develop. Frequent irrigation and fertilization in nurseries favors the development of pathogenic organisms, 

such as damping-off fungi. Well-processed, organic ingredients, especially composts, often discourage these harmful organisms.

Problems With Water and Air

Clay and silty soils have very small particle sizes and are therefore dense, heavy, and have few air spaces, making it difficult 

for water to drain freely out of the bottom of the container. Tropical soils tend to be rich in clay and low in organic material and 

are susceptible to compaction through routine handling and watering in the nursery. Heavy soils also accumulate at the bottom 

of containers, creating airless places inhospitable for root growth. In addition, some soil-based media shrink when dry and swell 

when wet, damaging plant roots.

Problems With Nutrition

In containers, native soils do not provide the quantities and ranges of nutrients needed for rapid plant growth and often 

immobilize certain nutrients (Landis 1995). Clays adsorb some nutrients so strongly that they become largely unavailable for 

plant uptake, while sandy soils hold nutrients poorly and lose most nutrients through leaching.

Variability and Weeds

Soils are naturally variable, so it is difficult to maintain the same quality from container to container, crop to crop, and year to 

year—making consistent crop production very difficult. In addition, soil also contains weed seeds, which compete for nutrients 

and water with the plants sharing their container and compel the grower to spend more labor on weeding.

Sustainability Concerns

Ecological sustainability must also be considered. Harvesting topsoil is actually a mining operation that uses up a limited 

resource that took thousands of years to develop. For example, in Mexico, millions of square meters of soil are mined each year for 

use in tree nurseries, leaving behind many acres of depleted areas where plants cannot grow well (Wightman 1999).

Because of these problems with field soil, growing media for tropical nurseries will ideally be based upon organic ingredients, 

such as composts, bark, rice hulls, or other materials, not field soil. If a medium containing soil must be used, it should contain 10 

to 30 percent soil at most, amended with organic ingredients to promote aeration and drainage while maintaining water-holding 

abilities (Landis 1995). More detail on how to use organic materials to create growing media, and how to incorporate some soil in 

growing mixes if necessary, is described later in this chapter.

106 Tropical Nursery Manual
two or three ingredients selected to provide certain physical, 

chemical, or biological properties. Mixtures of organic and 

inorganic ingredients are popular because these materials 

have opposite, yet complementary, properties (table 6.1).

Organic Ingredients

Common organic ingredients include compost, coconut coir, peat moss, bark, rice hulls, sawdust or any other 

appropriate, locally available material. These materials are 

lightweight, have high water-holding capacity and CEC, 

and some contain minor amounts of mineral nutrients. 

Some of these organic ingredients require screening or 

composting of local raw materials before use. The nursery 

may choose to do the processing, or a local supplier may 

specialize in the composting or processing local materials 

to sell to the nursery at a reasonable cost.

Peat Moss

Sphagnum peat moss is currently the most common 

organic component of growing media in temperate zone 

nurseries (figure 6.5A). Although types of peat moss may 

appear similar, they can have very different physical 

and chemical properties. The horticultural properties 

of Sphagnum peat moss (table 6.1) and the fact that it 

has uniform quality make it the only peat moss choice for 

plant nurseries that use peat moss. Most peat moss comes 

from Canada, some comes from New Zealand, and the 

one known tropical source is Indonesia (Miller and Jones 

1995). Therefore it is expensive and problematic to import 

peat for most tropical nurseries. In addition, extraction 

and transportation of peat moss on a large scale is a sustainability concern, and even temperate nurseries are 

considering alternatives. For tropical areas where peat 

moss is reasonably affordable and available, some nurseries use it in limited amounts for germinant mixes or for 

learning to grow unfamiliar native species. Some nurseries may use peat as a transition component, comparing 

peat’s properties to local materials such as composts or 

coir to develop local alternatives for growing media while 

moving forward with plant production.

Compost

Because of the risks of using soils and the expense of 

importing peat moss, many tropical nurseries prefer 

organic compost as a green alternative to peat moss (figure 6.5B). For example, in Florida, a variety of native plants 

grown in biosolid yard waste compost were as large or 

larger than those grown in a peat-based growing medium 

(Wilson and Stoffella 2006). Composts are an excellent sustainable organic component for any growing medium and 

significantly enhance the medium’s physical and chemical characteristics by improving water retention, aeration 

porosity, and fertility. Some composts have also been found 

to suppress seedborne and soilborne pathogens. Compost 

quality can vary considerably between different source 

materials and even from batch to batch so growers need to 

always test new materials before general use.

Composting is the physical and chemical decomposition of organic materials caused by the digestive activities 

Growing Media 107

Figure 6.5—Common organic ingredients of growing media include Sphagnum peat moss (A), compost (B), composted tree bark (C), 

coconut coir (D), and sawdust (E). Photos A and E by Thomas D. Landis, photo B by George Hernández, and photos C and D by Tara Luna.

B

C E

A D
108 Tropical Nursery Manual

Figure 6.6 —Creating good compost takes 1 to 12 months and requires the proper mix of organic materials and an ideal environment for 

the microorganisms that decompose the materials (A). Frequent mixing to foster good aeration is critical and can be done by hand (B) or using equipment (C). Compost goes through a typical temperature curve because of a succession of different microorganisms (D), so the process 

should be monitored with a long-stemmed thermometer (E). Illustrations A, B, and D from Dumroese and others (2008), photo C by Ronald 

Overton, and photo E by Thomas D. Landis.

B

E

A C

D
of insects, fungi, and bacteria (figure 6.6A). Raw materials for compost include any plant wastes such as vegetable 

or fruit scraps, leaves, weeds, or byproducts, such as cacao 

pods, coffee pulp, sugarcane bagasse, orchard prunings, 

and rice hulls; aquatic wastes such as aquatic weeds (such 

as the noxious weed water hyacinth) or fish parts from 

fish processing; animal wastes such as manures, feathers, 

and bedding; and wood wastes such as bark or sawdust. 

Sometimes these products are considered waste materials 

and are burned or disposed of at a cost to the producer—

composting turns them into a valuable resource. Organic 

nursery wastes may also be recycled through composting, including used growing media and culled seedlings. 

Sustainability and renewability of the compost source is 

important to consider. For example, seaweeds were once 

considered a good compost material, until people began to 

understand the importance of seaweeds to fish breeding 

and the damage that comes from depleting wild seaweed 

during fish-breeding season. Some nurseries grow part of 

their own compost ingredients using nitrogen-fixing trees 

or fast-growing plants such as comfrey, which can be cut 

back year after year. Growers anywhere should be able to 

find a sustainable source of organic matter that can be composted and used as a growing media component.

Composted tree bark from a wide variety of species has 

been successfully used as a growing media component (figure 6.5C), especially for larger volume containers. In the 

Southern United States, composted pine bark has become 

a standard ingredient in growing media for horticultural 

nurseries (Landis and Morgan 2009). The size of bark particles is important, and particle size can be controlled by 

hammer milling and screening (Gordon 2004). Obtaining 

bark of consistent quality can be a problem. Composting 

bark with supplemental nitrogen fertilizer supplies the 

nitrogen that microorganisms require during decomposition and helps lower the carbon-to-nitrogen (C:N) ratio.

Creating consistent-quality compost year after year is 

a challenging goal (Miller and Jones 1995). A nursery may 

choose to make its own compost or contract this work to 

a local processor. Ideally, compost is made from a mixture of organic materials. All material should be chopped, 

shredded, or cut into small pieces (0.5 to 2 in [1 to 5 cm]) 

to encourage faster decomposition and a more uniform 

final product. The most common method of making compost is to place organic materials in piles and allow them 

to decompose. The piles need to be mixed periodically to 

maintain adequate aeration and moisture needs to be maintained at about 50 percent with the feel of a damp sponge 

(figures 6.6B, 6.6C). Temperatures in the piles change over 

time as microbial decomposition progresses (figure 6.6D). 

Within the first few days, temperatures rise to 100 to 120 °F 

(38 to 49 °C) as the smaller and easily biodegradable materials decompose. Next, temperatures rise to 130 to 150 °F 

(54 to 65 °C) as more materials decay. A peak temperature 

of about 160 °F (71 °C) needs to be maintained for several 

days to kill weed seeds and fungal pathogens. Finally, temperatures fall to around 105 °F (40 °C) and lower during the 

“curing” stage. The process can be monitored with a thermometer (figure 6.6E). Mature compost can be produced in 

2 to 4 months in the humid tropics. It is important to note 

that the finished compost will only be about 40 percent by 

volume of the original fresh material. Often, growers have 

Growing Media 109

Figure 6.7—The maturity of commercial or homemade compost needs to be checked before use in growing media; in mature 

compost, original components are no longer visible and the material is dark and crumbly (A). Earthworms and soil insects are 

often visible in mature compost (B). Photo A by Tara Luna, and 

photo B by Thomas D. Landis.

B

A
two or more compost piles in varying stages of decomposition to have a continually available supply. For detailed 

guidelines on making compost, see Wightman (1999), 

Martin and Gershuny (1992), and Castillo (2004).

Mature compost should not produce an unpleasant odor 

or heat before incorporating into a growing medium. The 

compost should be dark in color and have a rich, earthy smell 

(figure 6.7A). The texture should be friable and crumbly; 

the original organic materials should not be recognizable. 

Earthworms and soil insects can live in mature composts and 

can be a sign that the compost is complete and ready to use 

(figure 6.7B). To determine if compost is ready, place two 

moist handfuls in a plastic bag, seal it, and leave in a dark, 

cool place. After 24 hours, open the bag: the compost is 

not ready if it feels hot or smells like manure or ammonia. 

Finished composts should be sifted through a screen similar to that used for field soil before use. Compost can be 

tested by sending to a soil lab, testing with a “bioassay,” 

and testing with an EC meter as described in the section 

on testing growing media.

C:N is a good indicator of whether nitrogen will be limiting or excessive (Landis and Morgan 2009). The higher 

C:N is, the higher the risk of nitrogen being unavailable 

to plants. The carbon in easily decomposed compounds, 

such as sugars and cellulose, are quickly used as an energy 

source by soil microorganisms, which also need nitrogen 

for growth and reproduction. Because this nitrogen is 

stored in their cells, it is unavailable for plant uptake. As 

carbon sources become depleted, the high populations of 

soil microorganisms gradually die and nitrogen is released 

for plant growth. When C:N is greater than 15:1, available 

nitrogen is immobilized. When C:N is below 15:1, however, 

nitrogen becomes available for plant uptake. Some composts have C:N as low as 10:1, indicating they are such a 

ready source of available nitrogen that they are considered 

fertilizers. Wood wastes, such as sawdust, have very high 

C:N (400:1 to 1,300:1). These materials are often composted with manure or supplemented with fertilizer to supply 

the needed nitrogen. The C:N of tree bark can be considerably lower than sawdust (70:1 to 500:1), and has become a 

preferred material for horticultural composts.

Coconut Coir

A byproduct of processing coconut husks is known as 

coir dust, coco peat, or simply coir. This material has proven 

to be an excellent organic component for container growing 

media and is readily available in some tropical locales (figure 6.5D). Coconut coir has many desirable qualities: high 

water-holding capacity; excellent drainage; absence of weeds 

and pathogens; physical resiliency (withstands compression 

of baling better than Sphagnum peat); slow decomposition; 

easy wettability; and acceptable levels of pH, cation exchange 

capacity, and electrical conductivity.

Coir is very similar to peat in appearance and structure, 

and, like peat, physical and chemical properties of coir can 

vary widely from source to source (Evans and others 1996; 

Noguera and others 2000). Coir is low in nitrogen, calcium, 

and magnesium but can be relatively high in phosphorus 

and potassium (Noguera and others 2000).

Excess salinity and phenolic compounds in coir can be a 

problem in areas with inadequate quality control (Ma and 

Nichols 2004). In addition, some coir sources have reportedly contained chlorides at levels toxic to many plants. 

Thus, it is very important that salts and other compounds 

are thoroughly leached with fresh water before shipment 

and use. Compared with Asia, little coir production occurs 

in tropical America, and, currently, supplies of coir are limited in some areas. Nurseries must locate a quality, consistent source and then add coir to media on a trial basis first, 

testing effects on a species-by-species basis.

Sawdust

Raw sawdust, with its high C:N, can negatively affect 

nutrient availability, especially nitrogen but its properties can 

be improved with composting (figure 6.5E; Miller and Jones 

1995). Also, because of inherent differences in chemical properties between different woods, the suitability of sawdust as 

an organic growing media component is extremely variable. 

Some species produce sawdust with phytotoxic effects. Only 

consider using sawdust from sawmills because other wood 

residues, such as from treated boards, may contain preservatives or harmful chemicals. Sawdust from coastal sawmills 

can contain high levels of salts, so all potential sources need to 

be tested before general use in the nursery.

Rice Hulls

Rice hulls are the sheaths of rice grains, a waste product 

of rice processing (Landis and Morgan 2009). Rice hulls or 

husks have been used as a component of potting medium 

with locally obtained peat for many years in Indonesia 

(Miller and Jones 1995). Several nurseries have used composted, screened, and hammer-milled rice hulls in place of 

composted bark (Landis and Morgan 2009).

Other Possible Organic Ingredients

Nearly any other organic material that is locally available has the potential to be an important addition to nursery 

growing media. Composted material takes longer to produce, 

but has a more reliable texture and nutrient content than raw 

material. For example, composted manure from livestock 

110 Tropical Nursery Manual
pens and other organic waste from agricultural operations 

are excellent candidates and are frequently available for free 

if you can haul them. All “homemade” materials will take 

effort to process and fine-tune to create a consistent product. 

The final product will be worth the effort because you will be 

developing your own specialized growing media with lowcost local ingredients that do not have to be shipped.

Inorganic Ingredients

Inorganic materials are added to growing media to produce and maintain a structural system of macropores that 

improves aeration and drainage (Mastalerz 1977). Many 

inorganic ingredients have a very low CEC and provide a 

chemically inert base for the growing medium. Inorganic 

materials with high bulk densities provide stability to large, 

freestanding containers. Several materials are routinely 

used as inorganic ingredients in growing media in native 

plant nurseries, including gravel, sand, vermiculite, perlite, 

pumice, and polystyrene beads.

Vermiculite

Vermiculite is a common component (figure 6.8A) and is 

a hydrated aluminum-iron-magnesium silicate material that 

has an accordion-like structure (figure 6.8B). Vermiculite has 

a very low bulk density and an extremely high water-holding 

capacity, approximately five times its weight. This material 

also has a neutral pH, a high CEC (table 6.1), and contains 

small amounts of potassium and magnesium. Vermiculite is 

produced in four grades based on particle size, which determines the relative proportion of aeration and water-holding 

porosity. Grades 2 and 3 are most commonly used in growing media; grade 2 is preferred when more aeration porosity is 

desired, whereas grade 3 produces more water-holding capacity. A 1:1 mixture of peat moss and coarse vermiculite is a 

common growing medium mix in many temperate nurseries.

Perlite

Perlite is a siliceous material of volcanic origin (figure 

6.8C). Perlite particles have a unique closed-cell structure so 

that water adheres only to their surface; they do not absorb 

water as peat moss or vermiculite do. Therefore, growing 

media containing perlite are well drained and lightweight. 

Perlite is also rigid and does not compress easily, which 

promotes good porosity. Because of the high temperatures 

at which it is processed, perlite is completely sterile. It is 

essentially infertile, has a minimal CEC, and has a neutral pH 

(table 6.1). Perlite is typically included to increase aeration, 

and commercial mixes contain no more than 10 to 30 percent perlite. Perlite grades are not standardized, but grades 

6, 8, or “propagation grade” are normally used in growing 

media. Perlite grades often contain a range of particle sizes, 

depending on the sieve sizes used during manufacturing. 

One safety concern is that perlite can contain considerable 

amounts of very fine dust that causes eye and lung irritation 

during mixing. Wetting the material while mixing and wearing dust masks and goggles can reduce this risk.

Growing Media 111

Figure 6.8—Common inorganic ingredients of growing media. Horticultural vermiculite particles (A) look like accordions (B) because of 

their expanded structure of parallel plates that allow vermiculite to absorb water and dissolved mineral nutrients like a sponge. Perlite particles have a closed-cell structure that prevents water absorption and improves aeration and drainage (C). The particles of pumice (D) and 

cinders (E) also improve aeration. Sand is also used as an inorganic component but may affect pH (F). Photos A, B, C, and D by Thomas D. 

Landis, and photos E and F by Tara Luna.

B

C E

F

A

D
Pumice and Cinder (Scoria)

Pumice (figure 6.8D) is a type of volcanic rock consisting of mostly silicon dioxide and aluminum oxide with small 

amounts of iron, calcium, magnesium, and sodium. The 

porous nature of pumice particles improves aeration porosity 

but also retains water within the pores. Pumice is the most 

durable of the inorganic ingredients and so resists compaction. Cinder (often called scoria) is another type of volcanic 

rock and a common growing media component in volcanic 

areas such as Hawai‘i, where growers may sift the cinder rocks 

to obtain the desired sizes for their containers (figure 6.8E).

Sand

Sand is one of the most readily available materials and 

is relatively inexpensive. The composition of sand varies 

widely. When considering if local sand is a suitable component, the type of sand and sand particle sizes must be 

considered. For example, some silty river sands with small 

particle size can have a serious negative effect on growing 

media by making them excessively heavy and not contributing to improved aeration or drainage.

Nurseries with access to siliceous (granite or schist 

derived) sands may be able to use local sand as an inorganic 

component. Sands derived from calcareous sources (such as 

coral or limestone) (figure 6.8F) are high in calcium carbonate 

(CaCO3), however, and can have dangerously high pH values. 

Growers can test sands by adding a drop of dilute acid or even 

strong vinegar—a fizzing reaction indicates the presence of 

CaCO3. It is better not to use coral-based soils or sands if at all 

possible but, if necessary, add lots of organic matter to help 

buffer the effects. Some plants grown in your nursery may be 

adapted to local calcareous soil conditions and may not suffer 

from the increased pH if the sand is used sparingly.

Sand is used to increase porosity, but small sand particles 

can lodge in existing pore spaces and reduce aeration and 

drainage. In general, sizes of 0.002 to 0.010 in (0.05 to 0.25 

mm) are too small and will block drainage holes and reduce 

aeration (Wilkerson 2011). Larger (medium to course) particles are more suited to increase porosity. The general recommendation is to wash sand (flushing out salt content if 

present) and sterilize or pasteurize it before incorporating 

it in the growing medium (Miller and Jones 1995). These 

techniques are described later in the chapter. Perhaps the 

more serious drawback of using sand in growing media 

is its weight, which causes problems with handling and 

increases the cost of shipping (Gordon 2004).

Polystyrene Beads or Flakes

Polystyrene is more commonly known by its trademarked name StyrofoamTM. Beads or flakes of polystyrene are 

a processing byproduct. Polystyrene increases aeration and 

drainage, decreases bulk density, and is highly resistant to 

decomposition (Wilkerson 2011). New polystyrene is unlikely 

to be a locally available material and many people are phasing 

out the use of polystyrene for sustainability concerns. It may 

be possible to recycle polystyrene and use pieces in growing 

media although it is not biodegradable and is often considered undesirable to outplant on project sites.

Developing a Growing Medium

It is likely that as many recipes exist for growing 

media as the number of nurseries; there are no global 

recommendations (Jaenicke 1999). Every nursery manager needs to be able to experiment and find suitable, 

local, affordable ingredients to create good growing 

media. Three general types of growing media are used in 

container nurseries (table 6.2):

1. Seed Propagation Media. For germinating seeds 

or establishing germinants (sprouting seeds), the 

medium must be sterile and have a finer texture to 

maintain high moisture around the germinating 

seeds.

2. Rooting Cuttings Media. Cuttings are rooted with 

frequent misting, so the growing medium must be 

very porous to prevent waterlogging and allow good 

aeration necessary for root formation.

3. Transplant Media. When smaller seedlings or rooted 

cuttings are transplanted into larger containers, the 

growing medium is typically coarser.

Because of the diverse characteristics of various growing 

media ingredients, a growing medium can be formulated 

with nearly any desired property. The physical, chemical, 

and biological properties of each growing medium strongly 

interact with nursery cultural practices, however, particularly irrigation, fertilization, and the type of container. 

When considering a new growing medium, first test it on 

a small scale with several different species and evaluate 

its suitability before making a major change to the whole 

crop. Information about testing is provided at the end of 

this chapter, and more details about proper ways to test are 

in Chapter 20, Discovering Ways to Improve Nursery Practices and Plant Quality.

Purchasing a Commercial Mix

A variety of commercial mixes are available that feature combinations of organic and inorganic ingredients 

described previously. To appeal to a wider market, many 

brands contain a wide variety of additional amendments 

including fertilizers, wetting agents, hydrophilic gels, and 

112 Tropical Nursery Manual
even beneficial microorganisms. Many amendments are 

formulated for crops other than tropical nursery plants and 

may do more harm than good. In particular, hydrogels can 

cause growing media to retain too much water and actually 

decrease aeration porosity when expanded. Always check 

the label to be sure of exactly what is in the mix.

Creating a Custom Mix

Many tropical plant growers prefer to mix their own 

custom growing media (figure 6.9). In addition to saving money, custom mixing is particularly useful in small 

nurseries where separate mixes are needed to meet propagation requirements of different species. A very porous 

and well-drained medium, for example, might be needed 

for plants from very dry habitats, but a second type is 

needed for coastal plants, and a third is needed for wetland species.

Standard commercial ingredients, such as peat and 

vermiculite, can be bought in bulk (by the pallet or container load to islands) to reduce costs. These ingredients have been used worldwide and may provide good, 

basic media ingredients during nursery start-up phases. 

Because of the steep learning curve with growing tropical plants and running a nursery, some managers may 

choose to import expensive but well-researched ingredients until they identify a consistent supply of local 

equivalents. Others may use small amounts of imported 

ingredients for seed propagation or cutting propagation, 

but use local ingredients for transplants.

Growing Media 113

Media type Properties Examples of media (by volume) Reference

Seed 

propagation

Maintains uniform moisture around germinating 

seeds (not too wet or too 

dry); no fertilizer; free 

from pests and diseases.

• 3 parts perlite to 1 part coarse vermiculite 

(for beach plants)

• 4 parts perlite to 1 part peat

• 3 parts small rinsed cinders to 1 part peat and 1 part perlite

Lilleeng-Rosenberger 

(2005)

• Fine, washed quartz sand [0.02 to 0.04 in (0.5 to 1 mm)] (100% sand 

will need frequent watering) Jaenicke (1999)

Rooting 

cuttings

Porous to prevent waterlogging and to allow 

good aeration for root 

formation; provide support 

for cuttings; free from 

diseases and weed seeds.

• 3 parts perlite to 1 part vermiculite

• 3 parts small rinsed cinders to 1 part peat and 1 part perlite

• 100% rinsed small cinder (but needs frequent misting)

Lilleeng-Rosenberger 

(2005)

• 100% washed quartz sand (2 mm) Jaenicke (1999)

• 1 part grit or fine gravel to 1 part washed 

sand to 1 part aged sawdust

• 1 part grit or fine gravel to 1 part aged sawdust

Longman (1998)

Transplant

Coarser; heavy enough to 

keep plants upright; may 

contain some nutrients; 

free from diseases and 

weed seeds.

• 1 part peat and 1 part vermiculite

• 2 parts cinder or perlite to 1 part 

well-decayed compost and 1 part peat

Lilleeng-Rosenberger 

(2005)

• 1 part coarse sand, 2 parts coconut coir, 1 part topsoil/duff

• 2 parts bagasse to 1 part rice hulls and 1 part alluvial soil

• 1 part well-composed grasses to 1 part rice hulls or pumice

• 3 parts composed bark to 1 part sand and 1 part shale

Miller and Jones 

(1995)

• 2 parts well-decayed compost to 2 parts sand and 1 part 

clay soil Wightman (1999)

• 3 parts coir to one part compost

• 30% composted rice hulls, 50% pine bark, and 20% sand Lovelace (2011)

Table 6.2—Example growing media for different nursery uses. The ideal growing medium will vary among nurseries, environments, 

and plant species.
Use of Field Soil

The use of field soil is not recommended for growing 

nursery plants. Circumstances may require, however, that 

some nurseries include some field soil in their media while 

more affordable and sustainable alternatives are being 

developed, so soil-based media is discussed here.

When selecting soil, use dark topsoil that has a high percentage of organic matter; lighter sandy loams are better 

than heavy clays. After collection, sift the soil through a 0.5 

in (12 mm) screen to remove debris and large objects such 

as rocks (figure 6.10). When using field soils, heat pasteurization (described later in this chapter) can eliminate fungal 

pathogens, insect pests, nematodes, and weeds.

Soil-based mixes are safest for transplant media when 

transplanting into larger containers, such as polybags or 

1 gallon pots. The properties of soil-based mixes make 

them unsuitable for smaller containers, and the risk of 

disease makes them unsuitable in media for germinating 

seeds or rooting cuttings. Soil should comprise no more 

than 10 to 20 percent of the transplant media by volume 

although some nurseries use up to 30 percent. The remaining ingredients, for example, bark, sawdust, and pumice, 

promote drainage and aeration while maintaining a high 

water-holding capacity (Landis 1995).

Use of Amendments

An amendment is a supplemental material that contributes less than 10 percent of the mixture, whereas an ingredient usually constitutes a larger percentage. A variety of 

materials may be added to growing media during the mixing 

process, including fertilizers, lime, surfactants, hydrogels, and 

mycorrhizal inoculum. Many of these materials may not be 

114 Tropical Nursery Manual

Figure 6.9—Many growers prefer to mix their own speciesspecific media using different ingredients (A). Growing media 

including local sand and compost mixed with some imported 

ingredients is used and sold at the native tree nursery at Reserva 

Natural, Cañón de San Cristóbal, Barranquitas, Puerto Rico (B). 

Photo A by Tara Luna, and photo B by Brian F. Daley.

Figure 6.10—If field soil must be used as an ingredient, it needs 

to be screened for debris and other large particles before use. Photo 

by Brian F. Daley.

B

A
desirable, however, and, in fact, may be detrimental to plant 

growth because they are formulated for other crops. If the 

decision is made to use amendments, uniform incorporation 

is important because plant roots have access to only a limited 

volume of growing media in the relatively small containers 

used in tropical plant nurseries. Uneven mixing of incorporated fertilizers is one of the major factors causing uneven 

growth in container nursery stock (Whitcomb 2003). If you 

decide to use any amendments, test them first on a small 

scale, using the techniques described in Chapter 20, Discovering Ways To Improve Nursery Practices and Plant Quality.

Dolomitic Limestone. Called “lime” in horticulture, 

it has traditionally been added to growing media to raise 

the pH and supply calcium for plant nutrition. Better ways 

of supplying calcium exist, so we do not recommend limestone amendments unless the nursery is growing plants 

that require a neutral or alkaline pH.

Starter Fertilizers. Some commercial media contain 

a small “starter dose” of soluble granular fertilizer. If fertigation (irrigation water containing liquid fertilizer) is 

not possible, then starter fertilizer may be a good idea to 

ensure that young, developing plants have quick access to 

mineral nutrients. Because fertilizer is a salt, incorporating 

larger quantities of soluble fertilizer is never recommended 

because of the high potential for salt toxicity.

Controlled-Release Fertilizers. Commercial growing 

media and custom mixes often include controlled-release 

fertilizers. It is important to know the fertilizer formulation 

and release rate. See Chapter 12, Plant Nutrition and Fertilization, for more discussion about these fertilizers. Keep in 

mind that, when used in warm, tropical climates, controlledrelease fertilizers will have much faster release rates than 

indicated on the label, which are targeted to temperate areas.

Surfactants. These chemical amendments, also known 

as “wetting agents,” break down the surface tension of water 

and increase the wettability of hydrophobic organic materials such as peat moss and pine bark. Some surfactants have 

been shown to adversely affect plant growth. Before using 

surfactants, be certain that a problem really exists and can 

potentially be solved by using the product. Ask other nurseries about their experiences with surfactants and perform 

small tests before using them operationally.

Hydrophilic Gels (“hydrogels”) are cross-linked polymers that absorb many times their own weight in water. 

They have been proposed as additives to increase the 

water-holding capacity of growing media. Several brands 

of growing media contain hydrogels, but no empirical evidence shows they improve plant growth. Because nursery 

crops are regularly irrigated, the use of hydrogels is rarely 

justified.

Mycorrhizal Inoculum. One method of inoculating 

native plants with beneficial mycorrhizal fungi is to incorporate inoculum into the growing medium at the time of mixing. As with all amendments, this practice needs to be tested 

first before adopting it on a large scale. See Chapter 13, Beneficial Microorganisms, for more discussion on this topic.

Rock Phosphate. Rock phosphate may be added to 

increase phosphorous availability. Excessive phosphorus, 

however, may hinder development of mycorrhizae and 

also interfere with the absorption of other mineral nutrients (Wilkerson 2011).

Other Amendments. Some growers add small amounts 

of other amendments to their mixes, such as worm castings, 

bone meal, kelp, guano, humic acid, compost tea, and others. 

Consider any amendment carefully as to what it will accomplish, if it is necessary, and if it may do more harm than good.

Treatment of Growing Media 

Ingredients

Some growing media ingredients may need to be treated 

before mixing to reduce potential damage to plants. These 

treatments may include removing salts, killing unwanted 

organisms, and sifting.

Flushing Out Salts

Flushing out salts may be necessary for materials such 

as coir, sand, sawdust from mills near the ocean, and composts with excessive soluble salt levels. Leaching with fresh 

water can effectively lower soluble salts below damaging 

levels (Carrion and others 2006, Landis and Morgan 2009). 

Because of its low salinity, rainwater is ideal for this process 

but any fresh water source will work if enough is applied. 

Check the electrical conductivity (EC) of the leachate from 

the ingredients or composts to verify that the salts have 

been removed (as described in the following section on 

testing growing media).

Pasteurizing Ingredients

Sterilization refers to the complete elimination of all living organisms whereas pasteurization targets pathogenic 

fungi and bacteria. For growing plants, completely sterile 

growing media is not desirable because many beneficial 

microorganisms normally found in growing media can 

actually be antagonistic to pathogens. Some commercial 

growing media are pasteurized to prevent the introduction 

of pests, weeds, and diseases into the nursery. If concerned, 

contact the supplier to find out if their media has been 

treated. If you mix your own media, common inorganic 

ingredients, such as vermiculite and perlite, are inherently 

sterile; organic ingredients are suspect. The heat from the 

Growing Media 115
composting process will kill pathogens and other pests in 

composts, but when using field soil, growers need to seriously consider pasteurization.

Heat pasteurization is the most common way of treating growing media. Several heat sources can be used for 

pasteurizing growing media: moist heat from steam, aerated 

steam, or boiling water; dry heat from flame, electric pasteurizers, or microwave ovens; and solar heat. Ingredients 

and media are pasteurized commercially with large, expensive equipment, but smaller pasteurizing equipment is available for nurseries (figures 6.11A, 6.11C) and some nurseries 

have developed their own pasteurization process using fire 

or solar heat (figure 6.11B). A practical technique would be 

to enclose small batches of media spread thin (no more than 

6 in [15 cm] deep) under black plastic tarps on an inclined 

table or on top of a tarp to expose it to maximum sunlight. 

Long-stemmed thermometers can be used to penetrate the 

tarp in several locations to ensure that temperatures stay in 

the recommended range of 140 to 177 °F (60 to 80 °C) for 30 

minutes. After treatment, the material can be dumped off the 

table or tarp into a clean wheelbarrow or mixing area, cooled, 

and used in the growing medium.

Regardless of the heating method, it is important to 

maintain the entire mass of growing medium at a uniform 

temperature that exceeds the thermal mortality threshold 

of the various nursery pests. Pests may vary in their inability to tolerate high temperatures (figure 6.12), but most can 

116 Tropical Nursery Manual

Figure 6.11—For nurseries making their own media, pasteurization with steam (A) or wood heat (B) is simple, effective, and can 

be accomplished with portable equipment. Electric soil sterilizers 

can heat the soil long enough to eliminate most weed seeds and 

pathogens (C). Photos A and B by Thomas D. Landis, and photo 

C by J.B. Friday.

Figure 6.12—Necessary temperatures for heat pasteurization 

vary depending on the target pest. Temperatures need to be held for 

30 minutes in the target range to be effective on that pest. Illustration 

adapted from Baker (1957) by Jim Marin.

B

C

A
be eliminated by heating the growing medium to 140 to 177 

°F (60 to 80 °C) for at least 30 minutes. Excessively high 

temperatures can eliminate beneficial soil organisms and 

produce toxic chemical compounds.

Sifting or Screening Ingredients

Some ingredients, such as soil, sand, and cinder, may 

require screening or sifting to achieve the desired particle 

size (figure 6.13). Excessively small or fine particles can clog 

container drainage holes and reduce aeration whereas excessively large particles can interfere with container filling, root 

development, and plant extraction. It may be necessary to sift 

twice, once with a small mesh to eliminate material larger 

than desired, and a second time with a larger mesh to remove 

material smaller than desired.

Mixing Growing Media

Whitcomb (2003) emphasized that improper media 

mixing is one of the major causes of variation in container 

plant quality. Mixing should be performed by diligent, 

experienced workers who will faithfully monitor the growing media quality. Creating a uniformly mixed growing 

medium that has not been compacted, contaminated, or 

compromised is the challenge and the goal.

Small batches of growing media ingredients can be mixed 

by hand. Measure out the ingredients by volume and mix 

together in a wheelbarrow or bucket (figure 6.14A). Workers can mix larger batches on any clean, hard surface using 

hand shovels. Pile the ingredients on top of one another and 

broadcast any amendments over the pile. Then work around 

the edge of the pile with a large scoop shovel, taking one 

shovel full of material at a time and turning it over onto the 

top of the pile (figure 6.14B). Make sure that all parts of the 

pile are mixed by gradually moving the location of the pile 

Growing Media 117

Figure 6.14—Nurseries that mix their own media can do so by 

hand (A) or by using the moving pile technique (B). To reduce 

labor, a cement mixer (C) can also be used but care must be taken 

to avoid overmixing and resultant damage to particle size and 

structure. Photo A by Diane L. Haase, and photos B and C by 

Thomas D. Landis.

Figure 6.13—Screening growing medium ingredients may be 

necessary to achieve the desired particle size. Photo by Thomas 

D. Landis. B

C

A
to one side. Some organic ingredients repel water when dry, 

so frequently misting the pile with water at regular intervals during mixing improves water absorption. Continue 

this procedure until samples from the pile appear to be well 

mixed. Do not compress or compact the mixture.

Managers of nurseries that regularly require larger 

quantities of custom growing media should consider purchasing a mixer. A cement mixer (figure 6.14C) works well 

as long as care is taken to avoid excessive mixing, which 

breaks down the size and texture of ingredients. Fragile 

materials, such as vermiculite and peat moss, are particularly vulnerable to overmixing, which can easily happen 

when mixers are run too long, are overfilled, or if the ingredients are too wet. Over-mixed media compacts easily during container filling, which leads to reduced aeration and 

waterlogging.

Safety Considerations

Workers need to follow certain precautions when handling growing media or its ingredients, including the time 

spent filling containers. Dust is the most common concern, 

so work areas need to be well ventilated and workers need 

to wear protective dust masks and safety glasses (figure 

6.15). Misting the growing media and work areas with 

water reduces dust.

Perlite dust is of particular concern because of potential 

for silicosis, an inflammation that occurs over time when 

dust that contains silica is inhaled into the lungs. Based on 

medical studies, however, no relationship exists between 

handling perlite and the development of silicosis (Schundler Company 2002). The use of commercial vermiculite 

horticulture products presents no significant asbestos 

exposure risk to commercial greenhouse or home horticulture users (Chatfield 2001). Nonetheless, dust is irritating 

and common sense dictates that proper safety precautions 

be taken.

Workers with cuts or abrasions on their hands need to 

be especially careful handling Sphagnum peat moss because 

of sporotrichosis, which is a condition caused by a fungal 

pathogen sometimes found in peat moss and other organic 

materials. The spores of this fungus can invade cuts on the 

hands or arms of workers or can even be inhaled (Padhye 

1995). Preventative measures include the following—

• Store peat moss and peat-based growing media under 

dry conditions.

• Ventilate work areas well.

• Wear gloves, dust masks, and long-sleeved shirts to 

protect hands and arms.

• Thoroughly wash arms and other exposed parts of the 

body with soap and water to reduce the risk of infection.

• Treat any injury that breaks the skin with a disinfectant, such as tincture of iodine.

• Regularly sweep and wash work areas.

Testing Growing Media

As each nursery develops its own growing media based 

on local ingredients, issues can arise because of variability 

in materials. Homemade materials, particularly composts, 

may vary in their quality despite best efforts to achieve a 

consistent product year after year. Purchased materials can 

also be variable on occasion depending on source and quality control procedures.

To preclude surprises, test compost and mixed growing 

media well in advance of use. It is also recommended to 

test and record the results of growing media batches that 

worked well, to compare them to new or experimental 

batches (Grubinger 2007), and to develop and refine suitable alternative mixes with similar favorable properties.

118 Tropical Nursery Manual

Figure 6.15—When mixing growing media or filling containers, 

nursery workers need to wear dust masks and safety glasses. Photo 

by J. Chris Hoag.
Plant Bioassay Test 

One easy and effective test is known as a plant bioassay (Grubinger 2007). Simply put a sample of the growing 

medium in the containers that will be used in the nursery, 

sow an abundantly available, fast-growing species into the 

medium, and observe how the planting performs over a 

few weeks. Some growers like to use more sensitive, slower 

growing crops to test their mixes. It is also a good idea to 

test a few seeds of other plant species intended for the mix 

in addition to the bioassay seeds. Do the seeds germinate 

as expected? As they grow, is damping-off observed? Are 

pest problems, such as gnats or maggots, emerging? Is the 

medium stable, not becoming compacted or waterlogged 

over time? If the results are not good, growers will be glad 

they tested the mix with readily available seeds instead of 

with rare native plant seeds. If the mix works, it is ready to 

try in the nursery.

Testing With an Electrical Conductivity 

(EC) Meter

The salinity (salt level) of the growing medium is a key 

parameter affecting the development and health of plant 

roots.  Salts may come from growing media ingredients, 

irrigation water, and from added fertilizers. Measuring EC 

is a way to measure the amount of nutrients and salts present to ensure they are in the appropriate ranges for the species grown. Excessively high salt levels can damage or even 

kill succulent young plants.

The best growers routinely measure the EC of their growing media ingredients, their growing media before and during a crop cycle, their fertilizer applications to ensure correct 

dosages, and their water quality. Some types of EC meters 

can only be used with aqueous solutions and so are ideal 

for measuring irrigation water and fertilizer solutions but 

are not ideal for growing media (for more details on proper 

technique with these meters, see Landis and Dumroese 

2006). Direct sensor models are small handheld devices 

and have probes small enough to be inserted directly into 

growing media (figure 6.16). The advantage of the direct 

sensor procedure is that readings can be taken quickly and 

nondestructively. To ensure the probe has good contact with 

the growing medium, always test at the same media moisture 

content. The recommendation is to monitor about 1 hour 

after irrigation or fertigation. Direct sensor testing works 

best with small containers; readings in larger containers can 

vary significantly so a couple of readings should be taken 

and averaged. Always take readings at a standard depth. If 

the probe is inserted into a medium containing controlledrelease fertilizers and the tip of the probe gets close to or 

punctures a prill, the EC reading might be extremely high, 

requiring a second insertion of the probe into a different 

area. Regardless of the type of EC meter used, it needs to be 

calibrated frequently to ensure correct readings.

Three things make using EC data challenging. First, EC 

can be measured in a variety of units, so pick a unit and 

stick with it. Conversions for the most common EC units 

are available in Landis and Dumroese (2006). Second, 

native plants vary considerably in their tolerance to salinity; a plant growing on the beach may have a much greater 

tolerance for high salt levels than a plant growing far inland 

or at higher elevations. Third, little information is available 

on the EC tolerances of many native plants. These challenges make providing an acceptable range for EC difficult, 

but start with general guidelines (table 6.3).

Regularly record EC values throughout each crop production cycle, especially during the Establishment Phase, 

and compare those with the quality of the final plants (see 

Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records). Often, it can be the change in EC values 

that is more important. For example, if during the course 

of 1 month, the crop growth is declining at the same time 

the EC values are dropping, this pattern indicates insufficient nutrients are available and fertilization needs to be 

increased. Conversely, if growth is declining and EC values are increasing dramatically, this pattern could indicate 

salt toxicity that can be remedied by reducing the fertilizer 

rates and leaching the medium with clear water. Collecting 

EC data for a couple of years will help any nursery manager 

hone in on the proper levels for their particular nursery 

and crops. More information about EC meters and water 

quality is in table 11.1 of Chapter 11, Water Quality and 

Irrigation.

Growing Media 119

Figure 6.16—Growing media salinity can be measured using an 

EC meter. Photo by J.B. Friday.
Sending Growing Media to a 

Soil-Testing Laboratory

For more formal testing, growing media samples can 

be sent to a soil-testing laboratory (private, local extension office, or land-grant university) for testing. A measurement of pH, soluble salts (electrical conductivity), 

and nutrients should be requested (Grubinger 2007). 

Results can vary among laboratories depending on their 

procedures, so it is best to stick with one lab for testing 

from year to year, provided that the data appears accurate and consistent.

Obtaining test results from laboratories may take a 

few days to several weeks, and bioassay results always 

take several weeks. Laboratory staff can help with interpretation of results. Results may indicate that the growing medium requires modification and further testing 

before it can be used. Therefore, start the testing well in 

advance of when the medium is needed. Changing to a 

new growing medium will also require adjusting irrigation, fertilization, and other cultural procedures, so other 

trials and experiments might be valuable (see Chapter 

20, Discovering Ways to Improve Nursery Practices and 

Plant Quality).

References

Baker, K.F. 1957. The U.C. system for producing healthy container-grown plants through the use of clean soil, clean stock, and 

sanitation. University of California, Division of Agricultural Sciences, Manual 23. Berkeley, CA: University of California. 332 p.

Buamscha, G.; Altland, J. 2005. Pumice and the Oregon nursery 

industry. Digger. 49(6): 18-27.

Bunt, A.C. 1988. Media and mixes for container grown plants. 

London, United Kingdom: Unwin Hyman. 309 p.

Carrion, C.; Abad, M.; Fornes, F.; Noguera, V.; Maquieira, A.; 

Puchades, R. 2006. Leaching of composts from agricultural wastes 

to prepare nursery potting media. Acta Horticulturae. 697: 117–124.

Castillo, J.V. 2004. Inoculating composted pine bark with beneficial organisms to make a disease suppressive compost for 

container production in Mexican forest nurseries. Native Plants 

Journal. 5(2): 181–185.

Chatfield, E.J. 2001. Review of sampling and analysis of consumer 

garden products that contain vermiculite. Hampshire, United 

Kingdom: The Vermiculite Association. http://www.vermiculite.

org/pdf/review-EPA744R00010.pdf. (February 2006).

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Evans, M.R.; Konduru, S.; Stamps, R.H. 1996. Source variation 

in physical and chemical properties of coconut coir dust. HortScience. 31: 965–967.

Gordon, I. 2004. Potting media constituents. International Plant 

Propagators’ Society, Combined Proceedings. 54: 78–84.

Grubinger, V. 2007. Potting mixes for organic growers. Brattleboro, VT: University of Vermont Extension. http://www.uvm.

edu/vtvegandberry/factsheets/pottingmix.html. (August 2011).

Jaenicke, H. 1999. Good tree nursery practices: practical guidelines 

for research nurseries. International Centre for Research in Agroforestry. Nairobi, Kenya: Majestic Printing Works. 93 p.

Johnson, P. 1968. Horticultural and agricultural uses of sawdust 

and soil amendments. National City, CA: Paul Johnson. 46 p.

Landis, T.D. 1995. Improving polybag culture for sustainable 

nurseries. Forest Nursery Notes. (July 1995): 6–7.

120 Tropical Nursery Manual

Table 6.3—Electrical Conductivity (EC) guidelines for artificial growing media. Adapted from Timmer and Parton (1982).

EC range (μS/cm) Salinity rating

0 to 1,200 Low

1,200 to 2,500 Normal

2,500 to 3,000 High

3,000 to 4,000 Excessive

Greater than 4,000 Lethal

μS/cm= microSiemens per centimeter
Landis, T.D.; Dumroese, R.K. 2006. Monitoring electrical conductivity in soils and growing media. Forest Nursery Notes. 

(Summer 2006): 6-10.

Landis, T.D.; Morgan, N. 2009. Growing media alternatives for 

forest and native plant nurseries. In: Dumroese, R.K.; Riley, 

L.E., tech. coords. National proceedings: forest and conservation 

nursery associations—2008. Proc. RMRS-p.58. Fort Collins, CO: 

U.S. Department of Agriculture, Forest Service, Rocky Mountain 

Research Station: 26–31.

Lilleeng-Rosenberger, K. 2005. Growing Hawaii’s native plants. 

Honolulu, HI; Mutual Publishing.

Longman, K.A. 1998. Growing good tropical trees for planting: 

volume 3, tropical trees, propagation and planting manuals. 

Edinburgh, United Kingdom: Commonwealth Science Council.

Lovelace, W. 2011. Personal communication (with Thomas 

D. Landis). Elsberry, MO. President and CEO, Forest Keeling 

Nursery, Inc.

Lovelace, W.; Kuczmarski, D. 1994. The use of composted rice 

hulls in rooting and potting media. International Plant Propagators’ Society, Combined Proceedings. 42: 449–450.

Ma, Y.; Nichols, D. 2004. Phytotoxicity and detoxification of fresh 

coir dust and coconut shell. Communications in Soil Science and 

Plant Analysis. 35:205-218.

Martin, D.L.; Gershuny, G. 1992. The Rodale book of composting. 

Emmaus, PA: Rodale Press. 278 p.

Mastalerz, J.W. 1977. The greenhouse environment. New York: 

John Wiley & Sons. 629 p.

Miller, J.H.; Jones, N. 1995. Organic and compost-based growing media for tree seedling nurseries. World Bank Tech. Pap. No. 

264, Forestry Series. Washington, DC: The World Bank. 75 p.

Newman, J. 2007. Core facts about coir. Greenhouse Management and Production. 27(2): 57.

Noguera, P.; Abad, M.; Noguers, V.; Puchades, R.; Maquieira, A. 

2000. Coconut coir waste: a new and environmentally friendly 

peat substitute. Acta Horticulture. 517: 279–286.

Padhye, A.A. 1995. Sporotrichosis—an occupational mycosis. In: 

Landis, T.D.; Cregg, B., tech. coords. National proceedings, forest 

and conservation nursery associations. Gen. Tech. Rep. PNWGTR–365. Portland, OR: U.S. Department of Agriculture, Forest 

Service, Pacific Northwest Research Station: 1–7.

Schundler Company. 2002. Perlite health issues: studies and 

effects. Edison, New Jersey: The Schundler Company http://www.

schundler.com/perlitehealth.htm. (February 2002).

Timmer, V.R.; Parton, W.J. 1982. Monitoring nutrient status of 

containerized seedlings. In: Proceedings, Ontario Ministry of 

Natural Resources Nurseryman’s Meeting, 1982 June, Thunder 

Bay, ON. Toronto, ON, Canada: Ontario Ministry of Natural 

Resources: 48–58.

Whitcomb, C.E. 2003. Plant production in containers II. Stillwater, OK: Lacebark Publications. 1,129 p.

Wightman, K.E. 1999. Good tree nursery practices: practical 

guidelines for community nurseries. International Centre for 

Research in Agroforestry. Nairobi, Kenya: Majestic Printing 

Works. 93 p.

Wilkerson, D. 2011. Texas greenhouse management handbook. 

College Station, TX: Texas A&M University, Texas Cooperative Extension, Extension Horticulture. http://aggie-horticulture.tamu.edu/greenhouse/nursery/guides/green/index.html. 

(August 2011).

Wilson, S.B.; Stoffella, P.J. 2006. Using compost for container 

production of ornamental wetland and flatwood species native to 

Florida. Native Plants Journal. 7: 293–300.

Additional Reading

Hundly, L. 2006. How to make compost: a composting guide. 

Dallas, TX: compostguide.com http://www.compostguide.com. 

(February 2006).

Kuepper, G.; Everett, K. 2010. Potting mixes for certified organic 

production. Horticultural Technical Note. Butte, MT: The 

National Center for Appropriate Technology (NCAT) Sustainable 

Agriculture Project. https://attra.ncat.org/attra-pub/viewhtml.

php?id=47. (August 2011).

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1989. 

The container tree nursery manual: volume 5, nursery pests and 

mycorrhizae. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 171 p.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1990. The 

container tree nursery manual: volume 2, containers and growing media. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 88 p.

Swanson, B.T. 1989. Critical physical properties of container 

media. American Nurseryman. 169 (11): 59–63.

Wilson, S.B.; Mecca, L.K.; Stoffella, P.J.; Graetz, D.A. 2004. Using 

compost for container production of ornamental hammock species native to Florida. Native Plants Journal. 4: 186–195.

Growing Media 121
122
123

A nursery container could be anything that holds growing media, drains, allows 

for healthy root development, does not disintegrate before outplanting, and allows for 

an intact, healthy root system to be removed with a minimum of disturbance to the 

plant. Understanding how container properties affect plant health and growth, as well 

as nursery operations, will help growers choose the best containers for their needs. 

Container type and dimensions affect root development, amount of water and mineral 

nutrients available for plant growth, nursery layout, bench size, production scheduling, and plant transportation method. After a container is selected, it can be expensive 

and time consuming to change to another type.

Most nurseries grow a wide variety of species and therefore several different containers are required (figure 7.1A). Container choice for a particular plant species 

depends on root system morphology, target plant criteria (see Chapter 3, Defining 

the Target Plant), and economics. In general, the following points hold true regarding 

container type:

• Plants that develop shallow, fibrous root systems, as most forbs do, grow better 

in shorter containers (figure 7.1B).

• Plants with long taproots, such as many kinds of trees, grow better in taller 

containers (figure 7.1C).

• Plants with multiple, thick, fleshy roots, and species with thick, fleshy rhizomes 

grow better in wide containers (figure 7.1D).

Containers

Thomas D. Landis, Tara Luna, and R. Kasten Dumroese 7

Facing Page: Nurseries growing a wide variety of species use many different container types. Photo of Po’o Wai U Nursery on the 

Big Island of Hawai‘i by Diane L. Haase.
In the ornamental trade, large individual containers 

are called “pots” or “cans,” but they are simply called 

“containers” in native plant nurseries. An inexpensive 

alternative to the plastic pot is the “bag” or polybag, 

which has the same general purpose and function. Plants 

grown in small-volume containers are often referred to 

as “plugs.” Plug seedlings are usually grown in individual 

containers called “cells” or “cavities” that are aggregated 

into “blocks,” “trays,” or “racks.” Often, many plants will 

be germinated in a tray, transferred to a small pot, and 

later transplanted to a larger container. Making the right 

container choices at the right time is an important task 

for a nursery manager. This chapter describes some of 

the key biological and operational considerations necessary for choosing appropriate container types.

Container Characteristics 

Affecting Plant Development

Volume

The volume of a container dictates how large a plant can be 

grown in it. Optimum container size is related to the species, 

target plant size, growing density, length of the growing season, and growing medium used. For example, to grow large 

woody plants for an outplanting site with vegetative competition, a nursery would choose large-volume containers with 

low growing densities. This method creates taller plants, with 

larger stem diameters, which have been shown to survive and 

grow better under these conditions.

In all nurseries, container size is an economic decision because production costs are a function of how many 

plants can be grown in a given space in a given time. Larger 

containers occupy more growing space and take longer to 

produce a firm root plug. Therefore, plants in larger con124 Tropical Nursery Manual

Figure 7.1—Nurseries use a variety of containers to produce different species and stocktypes (A). Some plants, including most forbs, 

grow best in shorter containers (B) whereas taprooted species do better in taller ones (C) and fleshy-rooted plants should be grown in 

short wide containers (D). Photo A by Diane L. Haase, and illustrations B, C, and D adapted from Dumroese and others (2008).

C

B

D

A
tainers are more expensive to produce, store, ship, and outplant. The benefits, however, may outweigh the costs if the 

outplanting objectives are more successfully satisfied.

Height

Container height is important because it determines the 

depth of the root plug, which may be a consideration on dry 

outplanting sites. Many clients want their plants to have a 

deep root system that can stay in contact with soil moisture throughout the growing season. Height is also important because it determines the proportion of freely draining growing medium within the container. When water is 

applied to a container filled with growing medium, gravity 

percolates it downward until it reaches the bottom and runs 

out of the drain holes in the bottom of the container. There, 

it stops because of its attraction for the growing medium, 

creating a saturated zone that always exists at the bottom 

of any container. Container height and the type of growing 

medium control the depth of this saturated layer. With the 

same growing medium, the depth of the saturation zone is 

always proportionally greater in shorter containers (figure 

7.2). For example, a 4-in (10-cm) tall container will have 

the same depth of saturation as a 10-in (25-cm) tall container, but the 4-in-tall container will have a smaller percentage of freely drained medium.

Diameter

Container diameter is important in relation to the 

type of species being grown. Broad-leaved trees, shrubs, 

and herbaceous plants need a larger container diameter so 

that irrigation water applied from above can penetrate the 

dense foliage and reach the medium. Diameter also affects 

growing density in the nursery.

Shape

Containers are available in a variety of shapes and most 

are tapered from top to bottom. Most containers are round 

but some are square and maximize the growing space 

used in the greenhouse. Container shape is important as it 

relates to the type of outplanting tools used and the type of 

root system of the species grown.

Plant Density

The distance between plants is another important factor to consider. Spacing affects the amount of light, water, 

and nutrients that are available to individual plants (figure 

7.3A). In general, plants grown at closer spacing grow taller 

and have smaller stem diameters than those grown further 

apart (figure 7.3B). Plant leaf size greatly affects growing 

density. Broad-leaved species should be grown only at fairly 

low densities, whereas smaller leaved and needle-leaved 

species can be grown at higher densities. Container spacing 

will affect height, stem straightness, stem diameter, and 

bushiness. Container spacing also affects nursery cultural 

practices, especially irrigation. Trays holding individual 

containers provide some flexibility in spacing because, as 

Containers 125

Figure 7.2—A saturated layer of growing medium always exists in 

the bottom of containers. With the same growing medium, the proportion of saturated media is greater for shorter containers. Adapted 

from Landis and others (1989).

Figure 7.3—Next to volume, spacing is the most important 

characteristic in container choice (A). Plants grown too close together become tall and spindly and have less stem diameter (B). 

Trays with removable containers are popular because they allow 

flexibility in spacing between plants(C). Adapted from Dumroese 

and others (2008).

B C

A
the plants grow, one-half of the containers can be moved 

to another tray, thereby allowing greater space between 

plants (figure 7.3.C). Some of the other effects of plant 

growing densities are shown in table 7.1.

Drainage Holes

Containers must have a bottom hole or holes large 

enough to promote good drainage and encourage “air 

pruning.” Roots stop growing when they reach an air layer 

under the container. Some containers feature a bottom 

rail to create this air layer (figure 7.4A), whereas flat-bottomed containers must be placed on specially designed 

benches (figures 7.4B, 7.4C) or on a bed of coarse gravel. 

The drainage hole must also be small enough to prevent 

excessive loss of growing medium during the containerfilling process. The correct drainage hole size is related to 

the type of medium you use; different types of containers work better with certain media types. Sandy mixes, 

for example, are more likely to fall out of large holes 

while media rich in fibrous organic material tend to hold 

together.

Root Pruning

Spiraling and other types of root deformation have been 

one of the biggest challenges for container growers, and 

nursery customers have concerns about potential problems 

with root-binding after outplanting (figure 7.5A). Research 

shows that rootbound seedlings are more likely to perform 

poorly or even blow over after outplanting. In addition to 

holes on the bottom of the container for air root pruning, 

many containers have vertical ribs within the container to 

force the roots to grow downward. 

126 Tropical Nursery Manual

Figure 7.4—Some block containers are designed to promote air pruning (A). Other containers must be placed on mesh-topped benches 

(B) or be supported (C) to create an effective air space underneath. Photo A by Thomas D. Landis, and photos B and C by Diane L. Haase.

Table 7.1—Effects of container density.

High density Low density

Plants will be taller and have smaller stem diameters Plants will be shorter and have larger stem diameters

Difficult to irrigate and fertilize with overhead sprinklers because water 

and liquid fertilizers need to penetrate dense patches of foliage Easier to irrigate and fertilize with overhead sprinklers

Greater likelihood of foliar diseases due to poor air circulation 

between plants Easier to irrigate and fertilize with overhead sprinklers

Cooler medium temperature Better air circulation between seedlings; less disease problems

Foliage in lower crown will die because of shading Plants have fuller crowns because more light reaches lower foliage

Greater number of plants can be grown in an area requiring less 

space and sometimes lower costs. Fewer plants can be grown in a given area of the nursery

A B C
Containers 127

Figure 7.5—Plants with aggressive roots often exhibit spiraling and other deformities after outplanting. If rootbound, roots often do not 

grow out beyond the original plug (A). Containers coated with copper will chemically prune roots (B), and other containers are available 

with lateral slits to reduce spiraling and encourage air pruning and on the side of the plug (C, D). Illustrations A, B, and C adapted from 

Dumroese and others (2008), and photo D by Thomas D. Landis.

Figure 7.6—Inappropriate containers and growing media can lead to serious problems, as with these 

mahogany trees in soil-filled 55-gallon drums. Photo 

by Brian F. Daley.

In an effort to save the cost of purchasing trees from the local 

nursery, an engineering firm decided to produce its own large 

mahogany trees for roadside planting projects. Because 55-gal 

drums were available in large quantities, they were cut in half and 

filled with local soil. Young mahogany seedlings were planted in 

the drums and watered regularly for nearly 3 years. The trees grew 

to be 15 to 20 ft (5 to 6.5 m) tall and were deemed to be the perfect 

size for the project (figure 7.6). The soil used was rich in clay and 

the containers did not have sufficient drainage holes, however, 

causing the root systems to be severely stunted. Even worse, the 

drums and clay made the trees too heavy to be transported easily. In the struggle to transport them and to remove them from 

the containers, the trees were damaged. In the end, transport costs 

tripled and one-third of the trees had to be replaced. Poor planning and improper containers turned this money-saving idea into 

a very costly project.

B C

A D

Inappropriate Containers 

and Growing Media
Chemical pruning involves coating the interior container 

walls with chemicals that inhibit root growth (figure 7.5B), 

such as cupric carbonate or copper oxychloride. Coppercoated containers are available commercially (such as the 

CopperblockTM). Some nurseries apply the chemical by spraying or dipping the containers. It can even be applied to polybags to greatly improve plant quality and outplanting performance, as described under the section on polybags. Fabrics 

treated with Spin-OutTM or similar copper coating can be 

placed as a groundcover under containers to prevent rooting 

into the ground. Copper toxicity has not been shown to be a 

problem for most native species, and the leaching of copper 

into the environment has been shown to be minimal.

Several companies have developed containers that feature air slits on their sides to control root deformation by air 

pruning (figures 7.5C, 7.5D). In the same manner as plant 

roots air prune when they reach the bottom drainage hole, 

they also stop growing and form suberized tips when they 

reach the lateral slits in sideslit containers. Using these containers (such as the RootMaker®) requires managing for (1) 

roots that sometimes bridge between containers and (2) 

seedlings in sideslit containers that dry out much faster than 

those in containers with solid walls.

Root Temperature

Color and insulating properties of the container affect 

medium temperature, which directly affects root growth. 

Black containers can quickly reach lethal temperatures in full 

sun whereas white ones are more reflective and less likely to 

have heat buildup. In hot, sunny climates, a grower should 

use containers in white or other light-reflecting colors to 

protect against root injury (figure 7.7). Another option is to 

use white plastic, StyrofoamTM, or other insulating material 

around the outside perimeter of the containers.

Economic and Operational Factors 

Affecting Container Choice

Cost and Availability

Associated expenses for various container types, such as 

shipping and storage costs, must be considered in addition 

to purchase price. Nursery managers in the tropics often 

face high shipping costs and long shipping times. Longterm availability must also be considered to ensure that 

ample supplies of the container can be secured in the foreseeable future. The environmental costs must also be considered and a number of new alternative container types 

are becoming available, made with recycled or green materials (Nambuthiri and others 2013).

Durability and Reusability

Containers must be durable enough to maintain structural integrity and contain root growth during the nursery 

period. Containers made out of rolled newspaper have gained 

popularity amongst gardeners; however, these containers 

disintegrate too quickly to be suitable for growing plants 

on a nursery scale. The intense heat and ultraviolet rays in 

container nurseries can cause even some types of plastics to 

become brittle and break, although many container plastics 

now contain ultraviolet (UV) inhibitors. Some containers are 

designed to be used only once, whereas others can be reused 

for 10 or more crop rotations. Reusability must be considered 

in the container cost analysis because the cost of reusable containers can be amortized over their life span after adjusting 

for the cost of handling, cleaning, and sterilizing of the containers between crops (discussed later in this chapter).

128 Tropical Nursery Manual

Figure 7.7—Container color is a consideration, especially when 

containers are exposed to direct sunlight. Roots in white containers 

stay cooler than those in black ones. Adapted from Dumroese and 

others (2008).

Refundable Deposits for 

Containers and Trays

Containers are expensive, and most are made of 

plastic—a nonrenewable, petroleum-based product. 

Although alternative containers made from renewable 

resources are being developed (Chappell and Knox 

2012, Nambuthiri and others 2013), many of these 

have not yet been adequately tested in tropical nurseries. Reusing containers is important to save money and 

resources and to protect the environment from waste. 

Charging a refundable container deposit (similar to 

bottle deposits for beverage containers) encourages 

clients to return containers to the nursery for reuse. 

Used containers need to be washed and sterilized after 

they are returned to the nursery.
Ease of Handling

Containers must be moved several times during crop 

production, so handling can be a major concern from logistic and safety standpoints. Collapsible or stackable containers, such as ZipsetTM Plant Bands or Spencer-Lemaire RootrainersTM, have lower shipping and storage costs; they must, 

however, be assembled before filling and sowing and thus 

require additional handling. The size and filled weight of a 

container will affect ease of handling. Containers must be 

sturdy enough to withstand repeated handling.

Large containers are increasing in popularity, but they 

become very heavy when saturated with water. Weight 

must also be considered for shipping and field planting. It 

is easier for a tree planter to carry and plant hundreds of 

trees if the seedlings are small. With large containers, the 

outplanting may go slower, but survival and ability to outcompete other vegetation will likely be greater. If containers will be shipped or trucked to the outplanting site, then 

the type of shipping and storage system must be considered 

during container selection. If seedlings are to remain in the 

container, then some sort of shipping box must be used to 

protect them. Pros and cons exist with every method, and 

it is up to you to know the needs of your plants, clients, and 

site conditions for outplanting to choose the best option, as 

discussed in Chapter 3, Defining the Target Plant.

Ability to Cull, Consolidate, and Space

One advantage of tray containers with interchangeable 

cells is that cells can be removed from the tray and replaced. 

This advantage is particularly useful during thinning, when 

empty cells can be replaced with cells containing a germinant, and during roguing, when diseased or otherwise undesirable plants can be replaced with cells containing healthy 

ones. Plants of the same size can be consolidated and grown 

under separate irrigation or fertilizer programs. This consolidation can save a considerable amount of growing space 

in the nursery. This practice is particularly valuable with 

seeds that germinate slowly or unevenly, and so exchangeable cells are very popular in tropical nurseries. Another 

unique advantage is that cells can be spaced farther apart 

by leaving empty slots; this practice is ideal for larger leaved 

plants and also for promoting good air circulation later in 

the season when foliar diseases can become a problem.

Holdover Stock

Some nurseries will hold onto their stock without transplanting in an effort to reduce costs and save growing space, 

hoping that the stock will be outplanted next year. This 

practice, however, can result in the root system becoming 

too rootbound to grow well after outplanting (figure 7.8), 

as discussed in Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records, and Chapter 15, Hardening. If nursery stock must be held over, then the nursery 

needs to keep a supply of larger containers so the plants 

can be transplanted to keep the root systems healthy and to 

maintain good shoot-to-root balance.

Types of Containers

Many types of containers are available, and each has its 

advantages and disadvantages. It is a good idea to try new 

containers for each species on a small scale before buying 

large quantities. Several containers types are used in container plant nurseries and can vary considerably in size 

(table 7.2).

One-Time-Use Containers

One of the first major distinctions in container types 

is whether they will be used once or whether they can be 

cleaned and used again. Some one-time-use containers, 

such as Jiffy® products, can be outplanted directly whereas 

others, such as ZipsetTM Plant Bands, are removed and discarded at the time of outplanting. The idea of growing a 

plant in a container that can be transplanted or outplanted 

directly into the field is attractive, and many designs have 

been tried. Most of these early attempts failed because the 

Containers 129

Figure 7.8—Many tropical plants have aggressive roots and cannot be held over from one growing season to the next or they will 

become dangerously rootbound. Photo by Thomas D. Landis.

“Seedlings produced in a nursery in Indonesia using 

root trainers and a peat-based growing media cost 

about 20 percent more than seedlings produced in a 

neighboring nursery using polybags and topsoil. But 

reduced transport costs to the field and better seedling 

survival and growth eliminated this difference completely.” (Jaenicke 1999: 27)
130 Tropical Nursery Manual

Container brand or type

Volume

 in3 

(ml)

Depth

 in 

(cm)

Top diameter

in

(cm)

One-time-use containers

Jiffy® pellets 0.6 to 21.4

(10 to 350)

1.2 to 5.9

(3 to 15)

0.8 to 2.2

(2.0 to 5.6)

Jiffy® pots 4.4 to 137.7

(72 to 2,257)

1.9 to 5.9

(4.9 to 15.0)

2.0 to 6.5

(5.1 to 16.5)

ZipsetTM plant bands 6 to 42

(98 to 688)

4 to 14

(10 to 36)

1.5 to 3

(3.8 to 7.6)

Single free-standing containers

RootMaker® singles 6.0 to 13.25

(172 to 1,125)

6.0 to 11.3

(15.3 to 28.7)

6.0 to 13.3

(15.3 to 33.8)

Polybags 90 to 930

(1,474 to 15,240)

4 to 8

(10 to 20)

6 to 8

(5 to 20)

TreepotsTM 101 to 1,848

(1,655 to 30,280)

9.5 to 24.0

(24 to 60)

3.8 to 11.0

(10 to 28)

Round pots 90 to 4,500

(1,474 to 73,740)

6 to 18

(15 to 45)

6 to 14

(15 to 35)

Containers with exchangeable cell held in a tray or rack

Ray Leach Cone-tainerTM cells 3 to 10

(49 to 164)

4.75 to 8.25

 (12 to 21)

1.0 to 1.5

(2.5 to 3.8)

DeepotsTM 13 to 60

(210 to 983)

3 to 14

(7.6 to 36)

2 to 2.5

(5 to 6.4)

Book or sleeve containers

RootrainersTM 3.3 to 67

(55 to 1,100)

4.25 to 10.0

(10.8 to 25.4)

1.0 x 1.0 to 3.0 x 2.5

(2.5 x 2.5 to 3.0 x 2.5)

Rectangular blocks composed of cavities or cells

StyroblockTM and CopperblockTM 0.5 to 195.3

(8 to 3,200)

2 to 7

(5.1 to 17.9)

0.6 to 6.2

(1.4 to 15.8)

Ropak® Multi-PotsTM 3.5 to 6.0

(57 to 98)

3.5 to 4.75

(9 to 12)

1.25 to 1.5

(3.2 to 3.8)

IPL Rigi-PotsTM 0.3 to 21.3

(5 to 350)

1.7 to 5.5

(4.4 to 14.0)

0.55 x 0.55 to 2.3 x 2.3

(1.4 x 1.4 to 5.9 x 5.9)

HikoTM Trays 0.8 to 32.3

(913 to 530)

1.9 to 7.9

(4.9 to 20.0)

1.0 to 1.3

(2.5 to 3.3)

Miniplug trays

“Groove Tube” SystemTM 1.4 to 11.7

(23 to 192)

2.50 to 5.25

(6.4 to 13.3)

1.13 to 2.25

(2.9 to 5.7)

RootMaker® propagation trays 1.6 to 6.0

(26 to 172)

2.0 to 4.0

(5.1 to 10.2)

6.0 to 13.3

(15.3 to 33.8)

Table 7.2—Volumes and dimensions of containers used in native plant nurseries. Adapted from Stuewe and Sons, Inc. (2013), and 

other sources.
container material broke down in the nursery before the 

plants were ready or they failed to decompose after outplanting. Still, new products are continually being introduced and are attractive to ecologically minded growers 

(Nambuthiri and others 2013; Chappell and Knox 2012).

Jiffy® Pellets and Pots

Jiffy® products consist of dry, compressed peat growing media inside a soft-walled, meshed bag and come in a 

variety of sizes (figure 7.9). When sown and irrigated, the 

pellet expands into a cylindrical plug surrounded by mesh 

that encourages air pruning all around the plug. Pellets are 

supported in hard plastic trays, so individual pellets can be 

consolidated to ensure full occupancy. Irrigation schedules 

must be adjusted (frequency increased) because of greater 

permeability of the container wall. Some root growth occurs 

between the pellets, so they must be vertically pruned 

before harvesting. Jiffy® Forestry Pellets are popular in forest nurseries in the Northeastern United States and Eastern 

Canada, where they are outplanted directly into the field. 

Smaller Jiffy® pellets are used for starting plants that are 

then transplanted into larger Jiffy® sizes or other containers. This system works well for species that germinate very 

slowly or over a long period of time.

ZipsetTM Plant Bands

ZipsetTM Plant Bands are square, one-use containers composed of bleached cardboard containers that are 

assembled in a hard plastic tray (figure 7.10). ZipsetTM 

Plant Bands maintain their integrity in the nursery but 

biodegrade after 9 to 18 months. Some tropical nurseries 

prefer ZipsetTM Plant Bands because they protect the root 

plug during storage and shipping.

Other Natural Fiber Containers

Containers made of fiber, such as coir or compressed 

paper, come in a variety of sizes (figure 7.11) and are popular with gardeners for vegetable seedlings. The roots can 

develop without the potential deformity problems of solidwalled containers, and natural fiber pots can be transplanted 

or outplanted with minimal root disturbance or transplant 

shock. When they are outplanted, roots penetrate the container as it breaks down. These containers have a few problems, however, when used in tropical nurseries. They may 

break down too quickly in warm, humid climates to be suitable for growing native plants. They tend to become coated 

Containers 131

Figure 7.9—Jiffy® pellets are composed of dry compressed peat 

surrounded by mesh and expand when watered. Smaller pellets 

are used to start germinants and can be transplanted into larger 

Jiffy® containers or other containers. Photo by Don Willis.

Figure 7.11—Recyclable containers like these constructed of 

paper fiber are ecologically friendly but do not hold up well in wet, 

humid climates such as greenhouses. Photo by Tara Luna.

Figure 7.10—ZipsetTM Plant Bands are inexpensive containers that can be shipped directly to the field. Photo courtesy of 

Stuewe and Sons, Inc.
with algae over time, which makes them slippery to handle 

and nearly impossible to store, so they present challenges 

for shipping and handling. If a nursery is growing some fastgrowing species that will not need to be transported very far 

before outplanting, natural fiber containers may be a suitable option. Natural fiber pots need to be tested before use to 

ensure they will not break down or become coated with algae 

too quickly for the growing cycle of the species propagated 

in them.

Single, Free-Standing Containers

Several types of single-cell containers are being used to 

grow native plants for specific conditions.

RootMaker® Containers

These unique containers have staggered walls and a staggered bottom that prevent root circling and direct roots 

toward the holes in the walls and the bottom of the container. 

The containers were among the first to use side “air slits” to air 

prune plant roots (figure 7.12) and are available in many sizes 

of single containers that are either square or round. Smaller 

volume RootMaker® cavities are joined together in blocks.

Polybags and Polytubes

Bags made of black polyethylene (poly) plastic sheeting 

are the most commonly used nursery containers in the world 

because they are inexpensive and easy to ship and store 

(figure 7.13A). It is unfortunate but polybags generally produce seedlings with poorly formed root systems that spiral 

around the sides and the bottoms of the smooth-walled con132 Tropical Nursery Manual

Figure 7.12—The RootMaker®, which was the first to feature sideslit air pruning, is available as a single, free-standing container or in 

aggregate blocks. Photo by Tara Luna.

Figure 7.13—Polybags are inexpensive containers (A) but can result in problems with seedling quality. Root spiraling is often serious, 

but polytubes in trays (B and C) and with copper-coating (D, on the right) can solve that problem. Photos A, B, and C by Tara Luna, and 

photo D by R. Kasten Dumroese.

A B C

D
tainers. This problem worsens when seedlings are held over 

and not outplanted or transplanted at the proper time.

In cases in which converting to hard plastic containers 

would be operationally or financially impractical, ways exist 

to improve container production using polybags. Some of 

these cultural modifications include (Landis 1995)—

• Managing container seedlings as a perishable commodity with a limited “shelf life.” This concept 

is particularly critical in tropical nurseries where 

seedlings grow year round. If seedlings cannot be 

outplanted when their roots fill the container, then 

they must be transplanted into a larger container. 

Holding over polybag seedlings is not an option.

• Using polytube containers (a polybag open at both 

ends, sometimes called a polysleeve) instead of polybags (figure 7.13B). These containers can usually be 

obtained from the same supplier as polybags or cut 

from a continuous roll with no bottom (Jaenicke 1999). 

Poly tubes eliminate much of the root spiraling. Poly 

tubes will hold growing media if they are properly 

filled and placed on elevated screen-bottomed trays to 

promote air pruning of roots (figure 7.13C).

• Using copper-coated polytubes or polybags. Plants 

grown in copper polybags produce a much finer, fibrous 

root system that is well distributed throughout the containers (figure 7.13D), but availability can be a problem.

• Switching from soil-based to “artificial” or organicbased growing media (composts, bark, or other 

materials instead of soil). See Chapter 6, Growing 

Media, for more information.

• Carefully transplanting germinants or direct-seeding 

into the polybag or polytube containers to avoid root 

deformations from improper transplanting of newly 

emerged seedlings.

TreepotsTM

These large-volume containers are constructed of flexible hard plastic and are good for growing trees and woody 

shrubs. Many sizes are available that are either square or 

round (figure 7.14A); square shapes increase space and 

irrigation efficiency in the growing area. TreepotsTM feature 

vertical ribs on the inside wall to prevent root spiraling, are 

reusable, and store easily because they can be nested when 

empty. The depth of their root plug helps plants access 

soil water on dry sites and, for riparian restoration, provides stability against water erosion. Because of their large 

height-to-diameter ratios, TreepotsTM require a support 

rack for growing and shipping (figure 7.14B), which can be 

made with fencing or other materials.

Round Pots

Round black plastic pots are available in many sizes 

from numerous manufacturers; one encouraging feature 

is that some brands are recyclable (figure 7.15A). Round 

pots are used in some tropical nurseries, especially for 

landscaping applications (figure 7.15B). Round pots 

are very durable and so can be reused for many years; 

because they can be nested when empty, they use little 

storage space. Most designs have a ridged lip that makes 

the pots easier to move and handle when they are wet. 

Root deformation has been a serious problem with these 

containers, although some are now available with internal ribs or copper coating to prevent root spiraling.

Containers 133

Figure 7.14—TreepotsTM are popular native plant containers because of their deep root systems (A), but they need to be held in a 

rack system (B). Photo A by Thomas D. Landis, and illustration B 

from Dumroese and others (2008).

B

A
Exchangeable Cells Held in 

a Tray or Rack

The major advantage of growing plants in individual 

cell containers supported in a hard plastic rack or tray is 

that the individual cells are interchangeable allowing for 

consolidation and spacing as described earlier. Racks are 

designed to create enough air space underneath to promote 

good air pruning. Plastic cells can be reused for several 

growing seasons.

Ray Leach Cone-tainerTM Cells

One of the oldest container designs on the market, the 

Ray Leach Cone-tainerTM cells are still popular with native 

plant growers. In this system, individual soft, flexible 

plastic cells are supported in a durable hard plastic tray 

(figure 7.16). Trays are partially vented to encourage air 

circulation between cells and have a life expectancy of 8 to 

10 years. Cells come in three types of plastic: recycled, low 

density, and low density with UV stabilizers. All have antispiral ribs and a center bottom drainage hole with three or 

four side-drain holes on the tapered end.

DeepotsTM

These single cells are constructed of thick plastic and held 

in hard plastic racks (figure 7.17). Available in several sizes, 

they have internal vertical ribs for root control and supports 

on the bottom of each container provide stability. Racks 

hold the containers together but do not create an air space 

underneath, so DeepotsTM must be grown on wire mesh or 

well-drained gravel to facilitate air pruning of the roots. Due 

to their large volume and depth, DeepotsTM are popular with 

native plant nurseries growing woody shrubs and trees.

134 Tropical Nursery Manual

Figure 7.16—Ray Leach Cone-tainerTM cells are one of the most 

popular container types for growing native plants (A) because 

they can be consolidated and spaced in the racks (B). Photos by 

Diane L. Haase.

Figure 7.15—Some standard round plastic containers or “cans” 

are now recyclable (A), and are sometimes used to grow native plants 

for ornamental landscaping (B). Photo A by Thomas D. Landis, and 

photo B by Diane L. Haase.

B B

A A
Book or Sleeve Containers: 

Spencer-Lemaire RootrainersTM

These unique “book” containers are composed of flexible plastic cells that are hinged at the bottom, allowing 

the growing media and root system to be examined during the entire growing season when the books are open 

(figure 7.18A). The books are held together in plastic 

or wire trays or “baskets” to form blocks of cells (figure 

7.18B). As the name implies, RootrainersTM have an internal rib system to guide plant roots to the drainage hole 

and to prevent spiraling. One real advantage of using the 

books is that they nest easily and can be shipped inexpensively; the nesting feature also makes for efficient storage. 

The plastic is less durable than other container types, but 

the books can be reused if handled properly.

Block Containers Made Up of 

Many Cavities or Cells

StyroblockTM and CopperblockTM

StyroblockTM containers are the most popular type of container used in forest nurseries in the Western United States 

and are available in a wide range of cavity sizes and densities 

(figure 7.19), although outside block dimensions are standard 

to conform to equipment handling. This container has also 

been used for growing native grasses, woody shrubs, and trees. 

The insulation value of StyrofoamTM protects tender roots from 

temperature extremes and the white color reflects sunlight, 

Containers 135

Figure 7.17—DeepotsTM come in large sizes and are popular for 

growing native shrubs and trees. Photo by Diane L. Haase.

Figure 7.18—Spencer-Lemaire RootrainersTM are designed to allow easy inspection of growing media and the root plug (A). The hinged 

soft plastic sheets are assembled and placed into hard plastic trays or wire “baskets” to form blocks (B). In addition, these containers 

encourage good root formation along the length of the plug (left) compared with spiraled roots formed in the bottom of a polybag (right) 

(C). Photos A and B by Thomas D. Landis, and photo C by J.B. Friday.

B

A C
keeping the growing medium cool. StyroblockTM containers are 

relatively lightweight yet durable, tolerate handling, and can 

be reused for 3 to 5 years or more. One major drawback is that 

plants cannot be separated and consolidated so empty cavities and cull seedlings reduce space use efficiency. Species with 

aggressive roots may penetrate the inside walls of the cavities 

(especially in older containers reused for several crops), making the plugs difficult to remove. The CopperblockTM container 

is identical to the StyroblockTM except that it is one of the few 

commercially available containers with copper-lined cavity 

walls to promote root pruning.

Hardwall Plastic Blocks

Hardwall plastic blocks are available in a variety of 

cavity sizes and shapes and outside block dimensions 

(table 7.2). Extremely durable, these containers have a 

life expectancy of more than 10 years. The thick plastic is 

impervious to root growth.

Ropak® Multi-Pots are white in color, available in 

square and round cavity shapes, and have been used to 

grow a wide variety of species (figure. 7.20A). Because 

they are so durable, they are popular in mechanized nurseries and have been used to grow herbaceous and woody 

species. Cavity walls have vertical ribs to prevent root spiraling. IPL® Rigi-PotsTM are usually black, but other colors 

can be obtained in large orders. They are available in a 

variety of block dimensions and cavity sizes and shapes 

including sideslit models to encourage air pruning of 

roots (figure 7.20B). The HikoTM Tray System features a 

variety of block and cavity sizes and shapes (table 7.2). 

All cavities have vertical root training ribs and sideslits 

(figure 7.20C). “The Groove Tube” Growing SystemTM features grooves in the side walls and large drainage holes to 

promote root development (figure 7.20D).

Miniplug Trays

Miniplug containers are used to start young seedlings 

that are transplanted into larger containers after establishment (figure 7.21). They are particularly useful for species 

with very small seeds that make precise seeding difficult. 

Multiple germinants can be thinned and plugs transplanted 

to larger containers. In these situations, the use of miniplug 

trays is much more space and labor efficient than direct 

seeding into larger cells. They require constant attention, 

however, because they dry out quickly. If miniplug trays are 

used, they may need to be irrigated several times a day or 

watered with automatic mist or subirrigation to keep them 

from drying out.

Seeds may also be started in trays, as discussed in 

Chapter 9, Seed Germination and Sowing Options.

136 Tropical Nursery Manual

Figure 7.19—A wide assortment of StyroblockTM containers is available (A). The cavity walls of CopperblockTM containers are coated 

with copper, which causes chemical root pruning of species with aggressive roots systems (B). StyroblockTM and CopperblockTM containers 

have been used to grow a variety of native plants. Photo A courtesy of Stuewe and Sons, Inc., and photo B by Thomas D. Landis.

A B
Cleaning Reusable Containers

Most nursery containers are reusable. Charging a deposit 

(for example 10 cents per cell, refundable if returned within 

30 days of plant delivery) or otherwise encouraging clients to return containers saves money and resources. 

Reusable containers usually have some residual growing 

medium or pieces of roots that could contain pathogenic 

fungi. Seedling roots sometimes grow into the pores of 

containers with rough-textured walls, such as StyroblockTM containers, and remain after the seedling plug 

has been extracted (figure 7.22A). Liverworts, moss, 

and algae also grow on containers and are very difficult 

to remove from reusable containers. Used containers 

Containers 137

Figure 7.21—Miniplug containers are used to grow small 

seedlings that are transplanted into larger containers. Photo by 

Tara Luna.

Figure 7.20—Hardwall plastic blocks are extremely durable containers made by several manufacturers. Ropak® Multi-Pots (A), 

IPL® Rigi-Pots (B), the HikoTM Tray System (C), and the “Groove Tube” Growing SystemTM(D). Photos A, C, and D courtesy of Stuewe 

and Sons, Inc., and photo B by Thomas D. Landis.

C D

A B
should be washed first to remove old growing media and 

other debris. If available, a pressure washer is excellent 

for this purpose. Otherwise, a regular hose, or a soak in 

a garbage can followed by a rinse with a hose, works too.

Next, the containers should be sterilized. Because many 

tropical nurseries choose not to use pesticides and chemical disinfectants, hot-water dips are the most safe, environmentally friendly, and effective way to kill fungi and other 

pests in used containers. Most pathogens and weed seeds 

are killed when containers are held at 158 to 176 °F (40 to 

60 °C) for more than 3 minutes (figure 7.22B). A good rule 

of thumb is to use a soaking temperature of 165 to 185 °F 

(75 to 85 °C) for 30 to 90 seconds for StyrofoamTM containers; 15 to 30 seconds is probably sufficient for hard plastic containers (Dumroese and others 2002). Soaking StyrofoamTM at hotter temperatures can cause the material to 

distort. Commercial units are available, but many nurseries 

have built homemade container dipping systems that hold 

the containers under hot water in a dip tank.

Other options for sterilizing containers and nursery 

tools include using household bleach or alcohol. These 

chemicals are phytotoxic to plants and should never be 

used on or near seedlings. For bleach, use regular household 

bleach (5.25-percent sodium hypochlorite concentration), 

diluted 1 part bleach to 9 parts water. Dip or soak the containers and tools in this bleach solution; then rinse them well 

before using. If the diluted bleach is stored in a covered container, such as a garbage can with a good lid, the solution 

may be used again, as long as it has not lost its strength. 

Wood, grain, or rubbing alcohol (70 to 100 percent) may 

also be used to sterilize containers and tools. Alcohol is 

used full strength. Dip or soak the containers and tools in 

the alcohol, but do not rinse them. Allow the containers 

and tools to dry in the air before using them.

References

Baker, K.F. 1957. The U.C. system for producing healthy container-grown plants through the use of clean soil, clean stock, and 

sanitation. University of California, Division of Agricultural Sciences, Manual 23. Berkeley, CA: University of California. 332 p.

Chappell, M.; Knox, G.W. 2012. Alternatives to petroleum-based 

containers for the nursery industry. Bulletin 1407. Athens, GA: 

University of Georgia, Cooperative Extension. 4 p. http://www.

caes.uga.edu/applications/publications/files/pdf/B%201407_1.

PDF. (March 2013).

Dumroese, R.K.; James, R.L.; Wenny, D.L. 2002. Hot water and 

copper coatings in reused containers decrease inoculum of 

Fusarium and Cylindrocarpon and increase Douglas-fir seedling 

growth. HortScience. 37(6): 943–947. 

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Jaenicke, H. 1999. Good tree nursery practices: practical guidelines for research nurseries. International Centre for Research in 

Agroforestry. Nairobi, Kenya: Majestic Printing Works. 93 p.

138 Tropical Nursery Manual

Figure 7.22—Used containers should be washed and sterilized before reusing (A), because residual growing media and seedling roots 

can contain disease organisms. Submersion in water of 165 to 185 

°F (75 to 85 °C) for 30 to 90 seconds has been shown to be adequate 

for all types of containers (B). Photo A by Thomas D. Landis, and 

illustration B adapted from Baker (1957) by Jim Marin.

B

A
Landis, T.D. 1995. Improving polybag culture for sustainable 

nurseries. Forest Nursery Notes (July 1995): 6–7.

Nambuthiri, S.; Schnelle, R.; Fulcher, A.; Geneve, R.; Koeser, A.; 

Verlinden, S.; Conneway, R. 2013. Alternative containers for 

a sustainable greenhouse and nursery crop production. HortFact—6000. Lexington, KY: University of Kentucky Cooperative 

Extension Service, Horticulture Department. http://www.uky.

edu/Ag/Horticulture/alternativecontainers.pdf. (March 2013).

Stuewe & Sons, Inc. 2013. Tree seedling nursery container catalog. http:/www.stuewe.com. (February 2013).

Additional Reading

Dumroese, R.K.; Wenny, D.L. 1997. An assessment of ponderosa 

pine seedlings grown in copper-coated polybags. Tree Planters’ 

Notes. 48(3): 60–64.

Landis, T.D.; Tinus, R.W.; MacDonald, S.E.; Barnett, J.P. 1990. 

The container tree nursery manual: volume 2, containers and 

growing media. Agriculture Handbook 674. Washington, DC: 

U.S. Department of Agriculture, Forest Service. 88 p.

Containers 139
140
141

Nurseries that work to strengthen and expand the presence of tropical native species are concerned about fostering diverse, strong, and well-adapted populations. For 

many tropical plants, however, the natural diversity of wild populations has been 

depleted. Habitat loss has reduced the range and sheer numbers of plants. For plants 

with commercial value, unsustainable harvesting practices may have reduced the 

numbers of plants with desirable characteristics while leaving behind inferior plants. 

The process of depleting a population of the best genetic properties so that future 

populations are weaker than the original populations is called genetic degradation. 

Seed collection for plant propagation is an opportunity to reverse trends of genetic 

degradation and species loss. Nurseries have a key role in conserving the gene pool of 

native plants. This chapter covers some important principles for genetic diversity and 

seed source selection; it also includes practical procedures for collecting, processing, 

and storing seeds.

Collecting, Processing, and 

Storing Seeds

Tara Luna and Kim M. Wilkinson

8

Facing Page: Seeds are removed from fruit and cleaned on screens in a shaded outdoor location at the nursery of Reserva 

Natural, Cañón de San Cristóbal, Barranquitas, Puerto Rico. Photo by Brian F. Daley.
Understanding Genetic Diversity 

and Seed Collection Ethics

Before creating a strategy for collecting native plant 

seeds, it is important to understand some key points 

regarding genetics and collection ethics. Seed collection 

strategies must protect genetic diversity for the future 

both at the collection sites and in the places where the 

offspring will be planted. On the outplanting sites, good 

seed collection practices ensure that inbreeding will not 

become a problem (Withrow-Robinson and Johnson 

2006) and that plant populations will be genetically viable to survive and adapt to new stresses. For restoration 

and conservation projects, maintaining genetic diversity 

is a key part of project objectives and of the target plant 

requirements (see Chapter 3, Defining the Target Plant).

Collecting Locally Adapted Native 

Seeds

It is critical to identify suitable seed collection sites. As 

much as possible, collect seeds from a habitat similar in 

elevation, aspect, and soils to that of the outplanting site to 

ensure local genetic adaptation. In some parts of the world, 

plant geneticists have defined “seed zones” for native 

species to show practitioners where locally adapted seed 

sources can be found for different restoration project locations. It is rare to find these recommendations for native 

tropical species, however, so collectors just have to do their 

best. Working with locally adapted plant sources is important not only for the survival and health of the plants, but 

also for the native birds, insects, and animals that depend 

on the plants (figure 8.1).

In addition, it is important to be sure the population you 

plan to collect seed from is of wild origin, not planted by 

people (BLM/SOS 2011). For example, do not collect seeds 

of native species that were planted as street trees, in landscaping, or as part of a restoration planting after a disturbance, because the genetic diversity and origin is uncertain. 

(This guideline does not apply to nonnative cultural and 

traditional species; although diversity is still important, 

these plants may be selected cultivars.) More details about 

choosing a seed collection location are addressed later in 

this chapter.

Use Collection Methods That Ensure 

Genetic Diversity

Because reproductive strategies vary by species, no 

standard collecting procedure exists that will ensure 

genetic integrity for all species. The Seeds of Success 

seed-collecting program (part of the Native Plant Materials Development Program, led by the U.S. Bureau of Land 

Management) has developed a useful general protocol, 

which is used by many Federal agencies and partners for 

seed bank collections. Key seed collection practices for 

ensuring genetic diversity are summarized in table 8.1. 

142 Tropical Nursery Manual

Figure 8.1—To maximize genetic diversity, observe flowering 

throughout the season so early, middle, and late bloomers are 

represented. Different flowering and dispersal times may interact with different pollinators. Photo by Brian F. Daley.

Why Collect Seed From More 

Than 50 Individual Plants?

(adapted from BLM/SOS 2011)

Research shows that seed collection from more than 

50 individuals results in the greatest proportion of alleles 

(genetic codes specifying certain traits) present in the 

field population, while still being practical. Some studies 

show that at least one copy of 95 percent of the alleles 

occurring in the population at frequencies of greater than 

0.05 can be achieved by sampling from—

1. 30 randomly chosen individuals in a fully 

outbreeding sexual species, or

2. 59 randomly chosen individuals in a 

self-fertilizing species.

The reproductive biology of many tropical species 

has not been studied, and to capture rarer alleles would 

require an increased sample size. Therefore, collectors 

are advised to sample from a single population with 

individuals of the target species in excess of 50 individuals, and to look for populations with larger numbers 

of plants.
Communicate About Seed Collection

When planning a seed collection strategy, some of 

these following communications will need to take place:

• In all cases, be sure to obtain landowner permission to 

collect seeds.

• Before collection, be absolutely certain of the species 

identification. If in doubt, collect a specimen and get 

help making a positive identification.

• If any other seed collectors are found using the site, 

cooperate with them to share the collection work 

and ensure that the genetic conservation practices 

described in this chapter are followed.

• Use great care in labeling seed sources and make 

sure that you, and the people who work with you, 

do not accidentally mix seeds with those from 

another plant collection area. Outbreeding depression resulting from the mixing of genotypes can 

potentially harm the population, resulting in the 

reduction of fitness and adaptive variation.

• For rare plant propagation, contact the appropriate 

organization in your area to obtain a collection permit in advance. These programs have jurisdiction 

over rare species and are responsible for monitoring and protecting the rare plant populations. The 

guidelines given in this chapter do not apply to 

threatened and endangered species; check with the 

appropriate organization to find the correct guidelines for each species in that category.

Collecting, Processing, and Storing Seeds 143

Seed collection method Rationale

Assess the target population and confirm that a sufficient 

number of individual plants (more than 50) have seeds at 

natural dispersal stage.

Ensure that adequate genetic diversity can be sampled from the 

population and that the seeds are likely to be at maximum possible 

viability and longevity.

Monitor seed maturation and assess insect damage and empty seeds 

throughout the population before making the seed collection. Carefully 

examine a small, representative sample of seeds using a cut test and for 

smaller seeds, a hand lens.

Estimate the frequency of empty or damaged seeds and confirm that 

the majority of seeds are mature and fully formed. (Seed development 

can vary within and between populations of the same species.) 

Collect seed from more than 50 individuals. 

Maximize the genetic diversity present within a collection of seeds. To 

gather the greatest proportion of alleles (genetic codes specifying certain 

traits) present in the field population as possible. 

Collect from distant individuals to reduce the chance of 

collecting only close relatives. Collect equally and randomly across the 

population, maintaining a record of the number of individuals sampled.

Capture the widest possible genetic diversity from the plant population 

and to avoid only a few genotypes being propagated. 

For trees, collect seeds or fruits equally from all parts of the crown—

top, sides, and bottom. Gather from individual trees at least 150 ft (50 

m) distant from each other (Dawson and Were 1997).

Ensure genetic diversity in tree seed, as these parts of the tree may have 

been pollinated at different times from different pollinators. Distance 

recommendation is to avoid collecting from closely related individuals 

(Dawson and Were 1997).

Return to the site to collect seeds from a population throughout its 

dispersal season. Collections taken from the same population during 

a single collecting season may be combined into one collection; do not 

mix collections between locations or years. Note multiple dates of collections on the seed label. 

Maximize genetic diversity in the collection, capturing early, middle, 

and late bloomers. (These different dispersal times may also interact 

with different pollinators.)

Collect no more than 20% of the viable seed available at the time of 

collection.

Ensure that the sampled population is not over collected and is maintainable, so seeds that are needed for natural dispersal or for local fauna 

will remain available.

Clearly label all bags (inside and out) with the species, date collected, 

location of collection, and name of collector. 

Ensure that each collection is properly identified so seeds can be used 

appropriately in restoration efforts.

Table 8.1—Seed collection techniques to safeguard genetic diversity of native species. Adapted from USDI BLM/SOS (2011).
Protecting the Seed Collection Site

Collection strategies need to be planned to minimize 

negative effects on the site. Collectors should follow the 

following steps:

• Avoid soil disturbance and plant damage while collecting seeds.

• Be sure not to overharvest seeds, leaving too few 

behind for natural regeneration and wildlife needs.

• Avoid collecting from weed-infested areas and do not 

transport weeds into pristine habitats and rare plant 

localities.

• If possible, allow collection sites to rest for at least 

two growing seasons between collections. Keep in 

mind that longer rest periods may be needed for 

some species and locations.

Understanding Flowers and Seeds

The tropics support the highest degree of species diversity in the world. It has been estimated that more than 50,000 

woody tree and shrub species are in this region. This diversity 

is also expressed in a wide range of flowers and fruits. Seed 

collectors and growers need to be able to distinguish fruits 

and seeds of species that they are collecting to ensure collection of the right structure at the right stage of development.

Plants are classified according to whether they produce 

spores or seeds. Spore-bearing plants such as ferns produce 

clusters of spores on the undersides of leaves that may or 

may not be covered with a papery covering. Spores can 

be collected like seeds just before they disperse, but they 

require special growing conditions to develop into plants. 

Seed-bearing plants are classified into two groups based on 

their flower types: gymnosperms and angiosperms.

Gymnosperms

Gymnosperms do not bear true flowers and are considered more primitive than angiosperms. Instead, gymnosperms bear male and female cones on the same tree. Male 

cones typically develop on the tips of branches and fall 

off after pollen is shed. Female cones enlarge and become 

more visible following pollination and fertilization, and 

seeds are borne naked on the mature scales. Gymnosperm 

cones can be dehiscent, indehiscent, or fleshy. Cones that 

are fleshy (such as Cycas) or surrounded by a fleshy aril 

(such as Torreya) resemble berries and are handled and 

processed in the same way (figure 8.2A). Dehiscent cones 

have scales that open at maturity to release the seeds (figure 8.2B) whereas indehiscent cones rely on animals to 

pry them open and disperse the seeds. In both dehiscent 

and indehiscent cones the seeds are usually winged (figure 

8.2C). Fleshy cones resemble berries and their seeds lack 

wings. Gymnosperm seeds are composed of the embryo, 

the nutritive tissue, and the seed coat (figure 8.2D).

Angiosperms

Angiosperms bear true flowers, and seeds are enclosed in 

an ovary that develops and surrounds the seeds after fertilization. Pollen is transferred from anthers (male reproductive 

144 Tropical Nursery Manual

Figure 8.2—Cones of gymnosperms can be fleshy or woody at maturity, as seen in Cycas (A) and Pinus (B) which release winged 

seeds (B, C). Gymnosperm seeds are composed of the embryo, the nutritive tissue, and the seed coat (D). Photos A, B, and C by 

Thomas D. Landis, and illustration D from Dumroese and others (1998).

B

C

A

D
Figure 8.3—Angiosperm fruits and seeds vary greatly among 

species; Calophyllum (A) and Minquartia (B). In some species, 

the embryo is not visible to the naked eye. Illustrations from 

Vozzo (2002).

structure) to the stigma surmounting the pistil (female reproductive structure). Following pollination and fertilization, the 

ovary enlarges into a fruit that contains one to many seeds. 

The fruit protects the seeds, provides them with nutrition 

during development, and helps with the dispersal of mature 

seeds. The seed is a ripened ovule consisting of a seed coat, 

the nutritive tissue (endosperm), and the embryo (figure 8.3). 

Embryo size varies widely among species.

Most angiosperms have perfect (bisexual) flowers, 

meaning they contain both the male and female reproductive structures in the same flower (figure 8.4A). Perfect 

flowers can be showy or very small and inconspicuous. 

Some species such as Swietenia and Ricinus have imperfect 

flowers, meaning that separate male and female flowers 

are borne in single sex flower clusters on the same plant 

(figures 8.4B, 8.4C). Some species are dioecious, such as 

Dondonea, which means that individual plants are either 

male or female (figures 8.4D, 8.4E). Thus, often only the 

female plants will bear fruits and seeds (figure 8.4F).

Because of the wide variety of flower types, resulting 

fruits also vary tremendously. Dry, dehiscent fruits are 

those that are woody or papery and split open at maturity. 

Some examples include capsules (figures 8.5A, 8.5B), some 

legumes or pods (figure 8.5C), and follicles (figure 8.5D). 

Dry, indehiscent fruits are those in which both the fruit and 

seed form an integrated part of the dispersal unit and do not 

split open at maturity. The thin shells that surround the seeds 

of these species are fused with the outer layer of the fruit and 

are dispersed as single units that resemble a seed and often 

have winged appendages. Examples of dry indehiscent fruits 

include achenes (figure 8.6A), schizocarps (figure 8.6B), nuts 

(figure 8.6C), and samaras (figure 8.6D). Some pods and capsules do not open at maturity and are basically handled as 

indehiscent fruits (figure 8.6E). Fleshy fruits are those where 

the tissue of the ovary is strongly differentiated. The pericarp 

is the part of a fruit formed by ripening of the ovary wall. It is 

organized into three layers: the skin (exocarp), the typically 

fleshy middle (mesocarp), and the membranous or stony 

inner layer (endocarp). These layers may become skin-like 

and leathery, fleshy, or stringy during development. Fleshy 

fruits such as berries, drupes, and pomes are indehiscent. 

Berries contain a fleshy pericarp with many seeds (figure 

8.7A) while drupes have a tough stony endocarp (known as 

the stone or pit) that encloses only one seed (figure 8.7B). 

Furthermore, some fruits are known as aggregate fruits, as 

seen in Ficus, Annona, and Morinda, which grow in a cluster of multiple fruits developed from a single flower and bear 

one seed each (figure 8.7C).

Seed Longevity

Concerning longevity, seeds can generally be classified 

into four groups: viviparous, recalcitrant, intermediate, 

and orthodox. Viviparous seeds are those that germinate 

before they are dispersed from the plant. The most common examples are some species of mangroves such as 

Avicennia and Rhizophora (figure 8.8A) and some tropical legumes (figure 8.8B).

Recalcitrant seeds only retain viability for a few days, 

weeks, or months. Most tree species from the humid 

tropics, where rainfall is distributed on a relatively even 

basis throughout the year, have recalcitrant seeds (Vozzo 

2002). In general, large-seeded species that drop moist 

from perennial plants in moist habitats are most likely 

recalcitrant (Hong and Ellis 1996) (figure 8.8C). Recalcitrant species are usually sown in the nursery immediately 

after collection.

Intermediate seeds are those that can withstand some 

degree of desiccation and storage at low temperatures, but 

they cannot tolerate freezing. Papaya has been stored sucCollecting, Processing, and Storing Seeds 145

B

A
146 Tropical Nursery Manual

Figure 8.5—Dry dehiscent fruits such as capsules (Slonanea species) (A) and 

Swietenia (B), legumes or pods (Acacia) (C), and follicles (Roupala montana) (D). 

Photos from Vozzo (2002).

Figure 8.4—Examples of flowers: Perfect bisexual flower of Hawaiian kokia (Kokia drynarioides) (A). Imperfect male (B) and female 

(C) flowers can occur on the same monoecious plant, as with Artocarpus species; imperfect dioecious flowers borne on separate plants of 

Dodonaea viscosa: male flowers (D), female flowers (E), mature seed capsules on a female plant (F). Photo A by Tara Luna, photos B, C, 

D, and E by Gerald D. Carr, and photo F by C.H. Lamoureux, courtesy of Gerald D. Carr.

B

C E

F

A

D

A B C

D
Collecting, Processing, and Storing Seeds 147

Figure 8.6—Dry indehiscent fruits—such as achenes (Asteraceae) (A), schizocarps 

(Hura crepitans) (B), nuts (Quercus) (C), and samaras (Terminalia) (D)—are actually 

a single unit where the fruit wall is fused to the seed. Some pods containing free seeds can 

also be indehiscent, while others are late dehiscent (Dipteryx) (E). Photos A and C by Tara 

Luna, and photos B, D, and E from Vozzo (2002).

Figure 8.7—Berries contain numerous 

seeds per fruit (A) (Myrica species), while 

drupes usually contain one seed surrounded by a stony pit (B) (Citharexylum 

spinosum). Noni (Morinda citrifolia) 

is an example of an aggregate fruit (C). 

Photo A from Vozzo (2002), photo B by 

Brian F. Daley, and photo C by Thomas 

D. Landis.

B

E

A C D

B

C

A
cessfully under conditions of 50-percent relative humidity 

with seeds dried to 10-percent moisture content for 6 years 

without affecting viability (Vozzo 2002). Other species that 

have shown intermediate storage behavior include neem, 

cinnamon, citrus, and coffee.

Orthodox seeds store easily for long periods of time 

because they tolerate desiccation. Tropical species that occur 

in areas of strong wet-dry seasonal cycles or semiarid environments along the coast often have orthodox seeds (figure 

8.8D). In general, dry, hard seed coats and small seeds from 

dry, dehiscent fruits are most likely to be orthodox.

Collecting Seeds

Effective native seed collection involves a number of 

steps to ensure quality seeds are collected at the right stage. 

Proper seed collection requires the following practices:

• Locate populations of desired species before or during 

flowering.

• Investigate the viability of seeds after dispersal or 

maturation on a species-by-species basis.

• Monitor potential sites directly after flowering when 

fruits are becoming visible.

• Record the dates of flower, fruit, and cone formation. 

Cones are often a 2-year crop, so you can assess cone 

crop the year before collection.

• Observe carefully the weather patterns during pollination, fruit formation, and maturation.

• Visit the site frequently to monitor the development 

and quality of the seed crop.

• Use collection dates from previous years to predict 

target collection dates and other information.

• Use a cutting test of a few sample seeds to determine 

maturity before collection.

• Collect seeds during dry weather, if possible.

Selecting Locally Adapted, Genetically 

Diverse Seed Sources

The genetics discussion at the beginning of this chapter 

detailed essential elements of seed collection to ensure local 

adaptedness and genetic diversity. These elements include 

collecting from a minimum of 50 individuals; collecting from 

distant individuals to reduce the risk of collecting only close 

relatives; collecting equally and randomly across the population; and collecting throughout the seed dispersal season 

(BLM/SOS 2011) (figures 8.9A, 8.9B). Choosing a seed 

collection site is crucial to ensuring that locally adapted, 

genetically appropriate materials are planted. Within each 

species are genetic variations. Local populations of native 

plants have adapted to local climate, soils, elevation, precipitation, environmental stresses such as wind or drought, and 

148 Tropical Nursery Manual

Figure 8.8—Viviparous seeds germinate and develop into a propagule before they disperse as seen in some mangroves (A), or while 

still enclosed in the fruit in some tropical legumes (B). Recalcitrant seeds, like palms (C), need to be collected before they germinate or 

lose their viability. Orthodox seeds, such as Leucaena species (D), typically have hard seed coats. Photo A by Thomas D. Landis, photo B 

from Vozzo (2002), and photos C and D by Tara Luna.

B

C

A D
other site conditions. Local native plant materials, collected 

from the same or similar habitats as the outplanting site, 

have been shown to perform better than nonlocal sources. 

Because seed zones have not been defined for most native 

tropical species, seed collectors must decide on a case-bycase basis what makes sense, considering climate, soil type, 

elevation, and other site conditions (Withrow-Robinson and 

Johnson 2006). Collectors should collect seeds from plants 

with high vigor and health. Collecting seeds from, and propagating, locally adapted plant sources improves not only 

survival and growth of the plants on the outplanting site, 

but also is important for the survival of native fauna, from 

insects to birds, that depend on these plants.

Selecting for Desired Characteristics

Tropical nurseries collect seeds for diverse clients 

and projects. Nursery-grown plants can be planted for many 

reasons: traditional uses, ornamental plantings, agroforestry, 

habitat restoration, silviculture, or a combination of these 

uses. Plants within a population can vary dramatically in their 

characteristics, such as the qualities, properties, and productivity of their fruit, wood, or medicinal products. For some 

projects, seed collectors may want to select for certain desired 

characteristics. Some of the desired characteristics (especially 

medicinal properties) may be difficult for seed collectors to 

discern. If in doubt, collectors can ask for help from end-users 

and clients to choose parent plants with preferred properties.

Seed selection may be for disease resistance. Depending 

on the disease vector, some plants showing obvious pest or 

disease problems may pass on that susceptibility to their 

offspring. Thriving local plants are excellent candidates 

as parents.

For plants with food, wood, fiber, or other uses, high 

productivity in terms of abundant fruit, nuts, foliage, or fast 

growth rate is often desired. Many native tropical species only 

produce large seed crops periodically. Heavy seed crops are 

often followed by light seed crops the following year. The 

interval between heavy seed bearing years is referred to as 

Collecting, Processing, and Storing Seeds 149

Figure 8.9—Good tree seed collection practices to maximize genetic diversity include collecting from individuals at least 150 feet (50 m) 

apart, collecting throughout the seed dispersal season, and collecting from throughout the canopy. Tools including pole pruners, ladders (A) 

and even equipment such as forklifts (B) are useful to help access the desired diversity. In addition to protecting genetic diversity, seed collection may involve selecting for certain desirable traits, such as the “canoe koa”: straight-boled, healthy, large trees (C). Photo A by Thomas D. 

Landis, photo B by Clark Allred, and photo C by Craig R. Elevitch.

B

C

A
periodicity. Depending on the goals of the project, collecting 

seeds across different periods may be advisable. For example, 

when restoring pollinator habitat or food for birds, continuity 

of supply even in sparse years is important.

For woody plants, growth form is also a key characteristic. For example, trees may range in form from small, multistemmed, shrubby individuals to large, straight-stemmed 

individuals. Depending on the preferred characteristics 

for the project needs, seed collectors can gather seeds from 

parents with the desired form. For example, in Hawai‘i, the 

renaissance of traditional Hawaiian canoe culture has regenerated interest in the koa tree, a species that can range from 

shrubby and branchy to tall and straight in form. “Canoe koa,” 

which are straight-boled, tall trees good for canoe making, are 

sought for some koa reforestation projects (figure 8.9C).

Choosing Quality Over Quantity When 

Collecting Seeds

Seed collecting is definitely an area where “quality over 

quantity” needs to be the standard to avoid perpetuating 

undesirable characteristics and eroding genetic diversity of 

the species. For example, sometimes plants produce high 

volumes of seed when they are stressed, diseased, or injured 

but those seeds may not be of high quality. Also, for tree 

seeds, it is much easier to collect from short, seedy, shrubby 

individuals than from tall, straight trees but collecting only 

from those easiest to reach results in less genetic diversity. 

For all plants, it is faster to collect from only a few individuals than to follow the recommendation to take small 

amounts from 50 or more individuals, but collecting with 

the goal of gathering the most seeds in the least amount of 

time is not an effective strategy. If working with contractors or nursery staff to collect seeds, it is recommended 

to pay collectors by the hour instead of by the pound, so 

that proper guidelines that ensure seed quality and protect 

genetic diversity are followed. The long-term ecological 

viability and future contribution of a planting is at stake.

Species Phenology

Monitoring species phenology (development throughout 

the season including flowering, fruiting and producing, and 

dropping leaves) is an important part of seed collection. Any 

observations may also provide clues on how to germinate the 

seeds. Good field experience has no substitute.

Be sure to record time and dates of flowering for each species. Flowering is easily observed in species with showy flowers but requires more attention for wind-pollinated species 

such as Podocarpus and Terminalia. Over time, recognizing 

the flowering sequence of the local flora enables staff to simplify the seed collection schedule by keying it to the flowering 

period of a few index species. Most developing fruits become 

visible only a few weeks after flowering and pollination. As 

you become familiar with the phenology of the species and 

local site conditions, you will be able to develop a seed collection schedule that is specific for your area.

Each species has its own flower and fruit arrangement, 

pollination strategy, and mode of seed dispersal. Some species will flower and fruit over an extended period of time 

while others will flower and fruit only once during a growing season. Different types of flower arrangements will 

have different blooming sequences. In tropical areas with a 

strong wet-dry seasonal cycle, most species will flower and 

fruit during defined periods. In tropical areas with an even 

distribution of rainfall, many species will flower and fruit 

irregularly throughout the year. These species produce several distinct fruit crops in 1 year within individual plants. 

For broad genetic representation, you will need to collect 

fruits from all periods of seed availability.

Fruits of many tropical species are available only for a 

short time after maturation. Some fleshy fruits can be picked 

before full maturation and ripened at the nursery without 

affecting seed viability. Other species will fail to ripen after 

they are detached from the tree. Collectors will also need to 

know how long the seeds of a species will remain viable on the 

ground. Collecting seeds from the ground needs to be timed 

correctly because a delay can result in loss of seed viability.

Often, a range of fruit maturity stages exists within a 

single plant. For example, some species have a flower stalk 

with a prolonged period of flowering and many different 

stages of fruit development. The seed collector will need to 

selectively harvest only the fully mature fruits and make 

repeated visits back to the collection site.

Be aware of the dispersal strategy of the species before 

attempting to collect seeds. Tropical fruits have developed 

many highly specialized strategies that aid in seed protection and seed dispersal away from the parent plant. Many 

species are dispersed by animals, including bats, monkeys, 

birds, and ants for which the fleshy fruits are food sources. 

Collectors need to time their collection before the fruit is 

consumed; in some cases, it may be necessary to bag or cage 

fruits to obtain seeds. Remember seed collection ethics and 

be sure to leave enough food for the native animals when 

collecting seeds.

Wind dispersal is very common for many tropical 

species. Seeds and fruits can have air-filled cavities, hairlike coverings, and various kinds of wings or parachutes 

(figure 8.10). Collectors need to time collection before 

seed dispersal and before windy days.

Some species such as Cassia and Mimosa disperse by force 

upon maturation. In these cases, you may need to bag the 

150 Tropical Nursery Manual
developing fruits with cloth to capture the seeds (figure 8.11). 

Use a fine mesh cloth with a weave that allows light transmission but is small enough to prevent the seeds from falling 

through the cloth. Tie the bags over developing seed stalks so 

seeds will be captured when they are dispersed by force.

Factors Affecting Seed Formation 

and Collection Timing

Environmental conditions can be either beneficial 

or detrimental to flowering and seed development. For 

example, drought and high temperatures may promote 

flowering, but prolonged moisture stress may cause plants 

to abort developing fruits and seeds or to have poor seed 

viability. Perform a cutting test on seeds directly before collection, as described in the following section.

Elevation, latitude, and aspect affect seed maturation 

because it is temperature dependent. Populations found 

on open, sun-facing slopes (south-facing north of the 

equator, north-facing south of the equator) will mature 

sooner than those on protected, shade-facing slopes. 

Low-elevation populations usually mature first, and seed 

collectors can follow seed maturity up slope with increasing elevation. Collectors should use favorable microenvironments to their advantage. For example, populations 

growing in full sunlight tend to produce more seeds than 

those that are heavily shaded. Plants growing on moist 

and nutrient rich soils will produce more and healthier 

seeds. Sites intensely browsed by livestock are poor 

choices for seed collection because animals often consume the current season’s growth, which limits flowering and seed production. Wildlife can quickly eliminate 

a maturing seed crop, and collectors may need to bag 

or cage developing fruits. Certain insects and fungi may 

also consume seeds. Fruits or seeds that have small exit 

holes, are discolored, or are misshapen must be avoided.

Ensuring That Seeds Are Healthy

The easiest way to ensure that seeds are healthy and 

ready for harvest is to use a cut test. A cut test allows 

you to inspect for mature, abnormal, infested, or empty 

seeds. Several seeds from several individuals within the 

population need to be examined. The two essential tools 

are a hand lens and a safety razor, knife, or scalpel for 

cutting. With care, cut the fruit or seed along its longest 

axis. Inspect the seeds for their internal coloring, how 

completely the internal tissue fills the seed coat cavity, 

and for the presence of an embryo. Depending on species, the embryo may completely fill the cavity or be tiny 

and embedded in the endosperm. (A microscope may be 

needed for examining very small-seeded species.) If the 

seed coat is soft and the contents are watery and soft, the 

seed is immature. If the seed coat is hard and the contents are firm and light tan to white in color, the seed 

Collecting, Processing, and Storing Seeds 151

Figure 8.10—Species that are wind dispersed must be collected 

as soon as they are mature. Shown: Heliocarpus appendiculatus. 

Photo from of Vozzo (2002).

Figure 8.11—Seeds that disseminate by force with explosive 

capsules are best collected by tying mesh bags over the developing fruits. Photo by Tara Luna.
is approaching maturity or is fully mature. Some species 

can be collected just before maturity if the entire inflorescence is cut and the seeds are allowed to cure properly 

before cleaning. In general, the optimum time for seed 

collection is when fruits are splitting open at the top.

Seed Collection Methods

The choice of seed collection method depends on the 

species to be collected. Some general collection methods include hand picking or hand stripping, collecting 

by hand from the ground after seeds have fallen, cutting 

fruit clusters, raking or shaking branches over a canvas tarp, bagging or caging developing fruits or cones, 

and tying canvas tarps between large woody plants. The 

following tools and supplies are useful when collecting 

seeds from natural stands:

• Labels, permanent markers, pencils, and seed collection forms to attach to bags.

• Scissors, pruning shears, and extendible pruning 

poles, or safety and tree climbing gear for taller trees.

• Hand lens to examine seeds to ensure they are full.

• Safety razor blades or sharp pocketknife for the cutting test of seeds and to examine fruits.

• Large paper bags for dry fruits.

• White plastic bags for fleshy fruits.

• Canvas tarps for collecting fruits from the ground.

• Hand gloves.

• Wooden trays for collecting seeds of low-growing 

plants.

• A storage box or cooler to keep collections from being 

overheated during transport.

• Binoculars for spotting fruits in taller trees.

• Fine mesh bags, cages, fine mesh cloth, and rubber 

bands for species with rapid dispersal.

Purchasing Seeds From a Reputable 

Source

Collecting yourself or contracting seed collection 

according to your specifications is the best way to ensure 

you get locally adapted, genetically diverse sources. Sometimes, however, you may need to buy seeds. If purchasing 

from a seed supplier without overseeing collection practices, 

it is best to ask what sources the company has available, 

rather than asking for a particular source—unscrupulous 

suppliers may claim to have exactly what you want. If you 

cannot locate a suitable source of seeds for purchase, it is 

best to collect your own. If your seed suppliers are willing 

and able to collect to your specifications, have them follow 

the guidelines provided in this chapter.

Purchased seeds must be of a high quality and free of 

weeds. When purchasing seeds, obtain and keep a certificate of the seed analysis for each seedlot. The seed analysis must have the scientific name of the species, cultivated 

variety (if applicable), origin of the seeds, an estimate of 

viability, the percentage of pure live seeds (PLS—discussed 

in the following section under Seed Testing), and the percentage of other crop seeds, weed seeds, and inert material. 

Purchase only seeds with high PLS values and very low percentages of weed seeds and other inert materials. It is often 

a good idea to ask about where the seeds were collected and 

to determine what weeds may be present in the seedlot.

Processing Seeds

The way in which seeds and fruits are handled during 

collection, temporary storage, postharvest handling, and 

cleaning can directly affect seed quality, viability, and storage life. Proper processing of fruits and seeds begins the 

moment the fruit or seed is removed from the parent plant. 

Proper processing includes short-term handling from the 

field back to the nursery, temporary storage at the nursery, and prompt and proper seed extraction if necessary. 

This step is followed by prompt and proper planting in the 

nursery (recalcitrant and orthodox seeds) or preparation 

for long-term storage (orthodox seeds).

In general, it is best to transport material from the field 

to the nursery as quickly as possible, avoiding exposure 

to direct sun, high temperatures, and physical abuse. Dry 

fruits, seeds, and cones can be left inside their paper collection bags for short durations. Placing plastic bags filled 

with fleshy fruits inside coolers will help prevent them 

from fermenting and being damaged by subsequent high 

temperatures.

Seed cleaning is necessary before sowing or long-term 

storage. In some cases, seeds will germinate slowly, or not at 

all, if they are not removed from the fruits.

Most tropical nurseries deal with small seedlots. Seed 

cleaning and processing can be laborious and time consuming, and specialized cleaning equipment can be expensive. A 

variety of inexpensive, low-tech methods and devices are easy 

to use, readily available, and work very well with a variety 

of fruit types. Some are described in the following sections. 

Whichever method of cleaning you choose, the seed cleaning 

area of the nursery needs to be well ventilated. Some fruits can 

cause allergic reactions, and fine dust can irritate skin, eyes, 

and lungs. It is important to wear gloves and dust masks during cleaning and to wash your hands afterward.

152 Tropical Nursery Manual
Recalcitrant Seeds

Recalcitrant seeds cannot withstand drying below a critical moisture level, so they are usually sown immediately after 

processing. During temporary storage before sowing, seeds 

must be kept fully hydrated by keeping them in trays under 

moist burlap or in plastic bags filled with moistened sand or 

peat moss in a shaded area with relatively cool temperatures. 

Relative humidity needs to be maintained at 80 to 90 percent.

Many species with recalcitrant seeds can be collected 

quite cleanly and are sown immediately without further 

cleaning. Others need additional cleaning that is typically 

accomplished by flotation in water. Immediately after collection, seeds are placed in a bucket of water. In general, the viable seeds sink whereas the nonviable seeds, trash, and debris 

float. As a side benefit, the soaking helps keep the seeds 

hydrated until they are sown. If seeds or fruits are collected 

from very dry ground, viable seeds may also temporarily 

float—a longer soak duration, perhaps even overnight, may 

be necessary to allow enough time for good seeds and fruits 

to hydrate and sink. Do a cut test to fine-tune this procedure.

Dry Fruits and Cones With 

Intermediate or Orthodox Seeds

After they arrive at the nursery, small quantities of dry 

fruits and cones can be dried in paper bags or envelopes as 

long as the contents are loose. Large quantities must be dried 

immediately by spreading the material evenly on a tarp or 

drying rack. A drying rack consisting of a simple wooden 

frame with multiple screens can be constructed at low cost 

and will make efficient use of space in a seed-drying room 

or greenhouse (figure 8.12). Drying racks can be made with 

mesh screens having fine holes that allow air movement but 

prevent seed loss. Different mesh screens will be necessary for 

different seed sizes. The materials will need to be turned several times per day to prevent it from becoming too hot, drying 

unevenly, or becoming moldy. Dry, dehiscent fruits, should 

also be covered with a fine mesh cloth as well to prevent the 

loss of seeds after fruits open. Good air movement, low relative humidity, and temperatures between 65 to 80 °F (18 to 

27 °C) promote even drying and eliminate moisture buildup 

that can cause mold and damaging temperature. A ventilated 

greenhouse or storage shed works well for this purpose. Temperature control is very important; use a shade cloth to 

keep temperatures from rising too high. Avoid rewetting 

dry fruits after collection. Also, make sure to exclude animals from your seed-drying area.

Separating seeds from dry, dehiscent fruits is usually 

easy because the fruits split open at maturity. Shaking the 

fruit inside paper bags so that the seeds fall out will readily separate small lots of seeds and the woody capsules can 

then be removed from the bag. Modified kitchen blenders 

with rubber-coated blades are very useful for cleaning small 

lots of dry fruits (Thomas 2003). The ideal amount of dry 

fruit material to place in a blender varies with its size, but 

one-fourth to one-third of storage capacity of the blender 

works well (Scianna 2004).

Screening is the easiest way to separate extracted seeds 

from debris such as dry leaves, wings, and small pieces of 

dried fruits. Screens can be constructed of hardware cloth 

and wooden frames. Commercial screens are also available 

in a range of sizes (figure 8.13A). At least two screen sizes 

Collecting, Processing, and Storing Seeds 153

Figure 8.12—Orthodox seeds require protection from wind during drying (A) and good ventilation to prevent mold development during post harvest handling and drying (B and C). Photos by Thomas D. Landis.

A B C
are needed. The top screen has openings large enough to 

allow the fruits and seeds to pass through, and the bottom 

screen has smaller openings that allow fine chaff, but not 

the seeds, to pass through. By placing the collected material 

on the top screen and shaking, most of the debris can be 

removed. When separating other small-seeded plants, such 

as sedges, rushes, and other tropical herbaceous species, 

you will need screens with very fine mesh or kitchen sieves 

to properly separate seeds from other debris (figure 8.13B).

If you are cleaning large, tough, leathery pods, you may 

need pliers, hammers, vices, or screwdrivers. Seeds that 

are contained in tough woody capsules can be extracted by 

heating the capsules in ovens or exposing them to fire in a 

portable barbecue grill. The heat treatment causes the pods 

to become brittle, which aids in the extraction of the seeds 

using hand tools. When using fire or heat and hand tools, 

be certain to not damage the seeds. By simply immersing 

them in water to separate fine chaff and other impurities, 

species with hard seed coats can be cleaned.

Conifer cones, after the scales open, can be placed in 

sack and shaken by hand or tumbled in a wire cage to dislodge seeds from the cone scales. Serotinous cones, such 

as those found on some tropical pines, require exposure to 

heat before the scales open. Cones need to be exposed to 

170° F (77° C) temperatures by placing the cones in ovens 

for a period of a few minutes to a few hours or by dipping 

them in hot water for a few minutes. If an oven is used, 

cones will need to be checked frequently during drying 

and removed when most have opened enough to allow the 

extraction of seeds. If the cones are dipped in hot water, 

the combination of heat and drying after the soak needs 

to be sufficient to open them. Most tropical conifer seeds 

are dewinged before sowing, which can be done by filling 

a burlap or cloth sack one-fourth full, tying or folding it 

shut, and gently kneading the seeds by squeezing and rubbing from outside the sack for a few minutes to detach the 

wings (figure 8.14). Repeat the screening process again with 

a mesh size that retains seeds but allows the smallest debris 

to pass through (Dumroese and others 1998). This method 

can be used on other winged tropical seeds as long as it does 

not damage the seeds. In cases where damage is a possibility, 

seeds are not dewinged and are planted as an intact unit.

The final step is fanning or winnowing, which separates 

detached wings, hollow seeds, and seed-sized impurities from 

good seeds (figure 8.15). When seeds are poured slowly in 

front of a small electric fan, they separate according to weight 

154 Tropical Nursery Manual

Figure 8.13—Screens are used to separate large debris and fine chaff from the seeds and are available commercially in a wide range of 

screen holes sizes to compliment any species (A). Seeds of many species are very small and require fine screens or sieves for cleaning (B). Photo 

A by Tara Luna, and photo B by Thomas D. Landis.

Figure 8.14—Conifer seeds are dewinged before sowing by 

gently kneading them in a cloth or burlap sack. Photo by Thomas 

D. Landis.

A B
from the base of the fan. Most heavy, sound seeds will come 

to rest near the base of the fan while hollow seeds, wings, 

and lighter impurities will tend to blow farther away. Moving from the fan outward, periodically collect a small sample of seeds and cut them in half to check for soundness, 

determining where the hollow seeds are and discarding 

them. All species will probably require several successive 

separations to obtain a desired degree of seed purity. A 

good target for most species is 90 percent or more sound 

seeds (Dumroese and others 1998).

Fleshy Fruits and Cones

Fleshy fruits and cones are very susceptible to fermentation, mummification, excessive heating, or microbial infestation, all of which can damage seeds. On the 

other hand, it is important not to let the fruits dry out 

because it can make cleaning them much more difficult. 

The best procedure is to temporarily store fleshy fruits in 

white plastic bags in a cool place or refrigerator until the 

seeds can be processed.

Seeds in fleshy fruits need to be processed shortly after 

collection. The first step in cleaning is to soak fleshy fruits 

in water to soften the pulp. The soak may need to last a few 

hours to a few days, depending on the species, and the water 

needs to be changed every few hours. After the pulp is soft, 

flesh can be removed by hand squeezing or mashing using 

a wooden block, rolling pin, or other device. The flesh can 

also be removed by wet screening, which involves handrubbing the fruits against screens using a steady stream of 

water to eliminate the pulp (figure 8.16). This method is 

used for most large fleshy tropical fruits. Another useful 

tool that can be used for small lots of fleshy fruits is the 

common kitchen blender or food processor with modified 

blades. Modified kitchen blenders can be used for small lots of 

fleshy and dry fruits after the impeller blades are coated with 

rubberized plastic coating (the material used to coat hand-tool 

handles) to prevent damage to the seeds (figure 8.17) (Thomas 

2003). Run them for about 1 minute to produce a puree of fruit 

and seeds. The puree should be placed in a bucket and water 

added slowly and continually resulting in most of the debris 

floating off and leaving clean seeds at the bottom of the bucket 

(Truscott 2004). Small, hobby-size rock tumblers (figure 

8.18) are also useful for cleaning small fleshy fruits or removing barbs or other appendages from seeds and fruits. Wet 

tumbling uses pea gravel or crushed stones and water in a 

Collecting, Processing, and Storing Seeds 155

Figure 8.15—Use a small fan to winnow empty seeds and wings 

from filled seeds. The heavier filled seeds will land closer to the fan 

while lighter, empty seeds will land farther away. Illustration from 

Dumroese and others (1998).

Figure 8.16—Easily clean fleshy fruits by using a simple mesh 

cloth and running water to remove pulp from seeds (A). Ripe fruit 

from coco plum or icaco (Chrysobalanus icaco), a coastal shrub native to Puerto Rico and the U.S. Virgin Islands, has the stringy flesh 

removed, leaving cleaned seeds ready for storage (B). Photo A by 

Thomas D. Landis, and photo B by Brian F. Daley.

B

A
rubber-lined tumbler vessel. Add only enough water to make a 

slurry from the gravel and fruit. The tumbler can be run overnight and checked the following day. After a course of tumbling, the contents are dumped into a sieve and the pulp or 

debris is washed off, leaving clean seeds (Dreesen 2004).

If fleshy fruits of species with dormant seeds are being 

cleaned, they need to be washed with water to remove any 

remaining pulp and dried for several days before storage.

Seed Testing

After seeds are cleaned, it is a good idea to determine 

their quality by testing seed viability, seed germination, or 

both. Seed viability tests estimate the potential for seeds to 

germinate and grow, whereas seed germination tests measure the actual germination percentage and rate. For seeds 

you are purchasing or selling commercially, you may also 

want to know the percentage of pure live seeds (PLS.)

Seed Viability Tests

Cutting tests, described previously, are the simplest seed 

viability tests and are usually performed during seed collection and often just before treating seeds for sowing. Cutting 

tests should also be performed on seedlots that have been 

stored for a long time to visually assess their condition. Cutting tests can reveal whether or not the seed is healthy, but 

really cannot determine anything about the potential for 

germination. A better test is the tetrazolium (TZ) test, a biochemical method in which seed viability is determined by 

a color change that occurs when particular seed enzymes 

react with a solution of triphenyltetrazolium chloride. Living tissue changes to red, while nonliving tissue remains 

uncolored (figure 8.19). The reaction takes place both with 

dormant and nondormant seeds, and results can be obtained 

within a couple of hours. Although the TZ test is easy to do, 

interpretation of results requires experience. For this reason, 

156 Tropical Nursery Manual

Figure 8.17—Modifying kitchen blender blades by using rubber 

coating allows the grower to clean small lots of dry and fleshy fruits 

without damaging the seeds. Photo by R. Kasten Dumroese.

Figure 8.18—A small hobby-size rock tumbler can be used to 

clean dry fruits with hard-to-remove appendages or fleshy fruits. 

Photos by Tara Luna.

Figure 8.19—Tetrazolium (TZ) tests stain living tissue red and 

can be used to estimate seed viability of a seedlot. Shown in this figure (left to right): dead embryo, damaged embryo, and healthy seed. 

From Stein and others (1986).

some larger nurseries or nurseries that also sell seeds send 

seed samples to seed analysts that have the necessary laboratory equipment and experience for testing. A third test is 

an excised embryo test. Embryos are carefully removed from 

seeds and allowed to grow independent of the seed tissue. 

Seeds often must be soaked for several days to remove hard 

seed coats, and excision of the embryo is an exacting procedure that normally requires the aid of a microscope. As when 

doing TZ testing, most nurseries send their seed samples to 

seed analysts for excised embryo testing.

Germination Tests

A seed germination test determines both germination rate 

and total germination percentage, and is used to determine 

sowing rates so seeds are used efficiently (figure 8.20). The 

germination rate indicates how promptly seeds germinate, 

whereas germination percentage indicates what proportion of 

the seeds eventually germinate. Germination tests are used to 

determine how many seeds to sow per container and how long 

you can expect seeds will continue to germinate after sowing. 

See Chapter 9, Seed Germination and Sowing Options, for 

details about sowing rates and methods.
If the species being tested has some type of seed dormancy, 

an appropriate treatment to remove dormancy will be needed 

before the germination test. Many nurseries will test dormant 

seedlots before and after the dormancy treatment to check 

its effectiveness. Actual germination in the nursery may vary 

greatly because of the inherent variability of germination in 

most plant species and differences in the environmental conditions during testing and growing at the nursery.

Use the following steps to conduct a germination test:

• Select an area in the greenhouse or office that can 

be kept clean.

• Line the bottom of plastic trays, Petri dishes, or 

similar containers with paper towels. For largeseeded species, line the bottom with moist sterile 

sand (bake sand in the oven at 212 °F [100 °C] 

for at least 1 hour to sterilize it) or unmilled 

Sphagnum moss.

• Moisten the paper towels or other substrate with 

distilled water.

• Remove equally sized seed samples from each container of the same seedlot, or, if only one container 

exists from the seedlot, remove the seeds from 

different portions of the container. Mix these 

samples together to form a representative sample 

(figure 8.21).

• From the sample, make 4 replicates of 100 seeds and 

spread each replicate onto the moist substrate in a 

container. The containers may be covered to reduce 

evaporation from the substrate.

• Use distilled water to remoisten the substrate as 

necessary, but never allow standing water to remain 

in the container.

• Place the containers under optimum germination 

conditions—ideally those in which light, temperature, and humidity can be controlled. Conditions 

similar to the nursery will yield more meaningful 

results.

• Count the number of germinants on a daily or weekly 

basis for up to 4 weeks on herbaceous species and up 

to 3 months on woody species. Uniformity of germination timing may be advantageous, but be sure you 

do not exclude healthy seeds that simply germinate 

more slowly from the genetic pool for that species.

Percentage of Pure Live Seed

The percentage of pure live seed (PLS) is a seed quality index that can be calculated during seed testing (figure 

8.22). When seeds are bought or sold, it is important to 

know the seeds have high PLS values and very low percentages of weed seeds and other inert materials. It is often a 

Collecting, Processing, and Storing Seeds 157

Figure 8.21—Test seeds by collecting primary samples from 

an entire seedlot to make up a composite sample. The composite sample is further divided into samples tested at the nursery 

or submitted to a seed laboratory for testing. Illustration from 

Dumroese and others (2008).

Figure 8.20—Paper towels (A), sterile sand (B), and Sphagnum (C) are suitable substrates for germination testing. Photos from 

Vozzo (2002).

A B C
good idea to ask about where the seed was collected and to 

determine what weeds may be present in the seedlot.

Storing Seeds

It can be quite beneficial to store seeds, especially for 

those species that yield seeds irregularly or to take advantage of a bumper crop of seeds. In addition, long-term seed 

storage is an important conservation method for threatened and endangered species. For proper seed storage, 

seeds must be mature and free of mechanical injury. The 

viability of seeds after storage depends on their viability at 

harvest, how they were handled during processing before 

storage, and storage conditions. Even under the best conditions, seeds degrade—the degree of longevity varies by 

species.

Storage Methods for Recalcitrant 

Seeds

As previously discussed, recalcitrant seeds of tropical 

species only retain viability for a short time; they are usually stored only temporarily before sowing. Some nutbearing species, however, can be stored for a few months 

as long as seeds retain high moisture content (35 to 50 

percent) under high relative humidity with good ventilation at cool temperatures. Recalcitrant seeds need to 

have constant gas exchange, so they are usually stored in 

unsealed containers in plastic bags filled with moist peat 

moss in a cool place, such as in the refrigerator. Ideal 

temperature for recalcitrant seed storage must be determined on a species-by-species basis.

Because recalcitrant-seeded species are numerous and 

ecologically important in the tropics, nursery propagation 

efforts play a key role in conserving and restoring tropical 

recalcitrant species (Kettle and others 2011).

Storage Methods for Intermediate and 

Orthodox Seeds

Intermediate seeds can be stored if the proper storage conditions are provided and seeds are dried to the appropriate 

level of seed moisture content. The ideal seed-moisture 

content and storage temperature varies among tropical 

species, however. In general, dried intermediate seeds cannot be stored below 50 °F (10 °C).

In nature, orthodox seeds of most tropical species occur 

in areas with a distinct wet-dry seasonal cycle, arid beach 

strands, savannahs, or high-elevation environments (as 

seen in Hawai‘i). For example, under proper storage conditions, the viability of Acacia koa seeds remained high 

after 25 years in storage (Young 1993). Orthodox seeds can 

be found in many tropical species within several families 

(Asteraceae, Brassicaceae, Chenopodiaceae, Combretaceae, 

Cucurbitaceae, Fabaceae, Lamiaceae, Poaceae, Rosaceae, 

Solanaceae, and Pinaceae). Species that produce achenes, 

many seeded berries, dry pods, follicles or capsules con158 Tropical Nursery Manual

Figure 8.22—Pure live seeds (PLS) is the percentage of the bulk seed weight that is composed of viable seeds. In this example, results of a 

purity test show 95 percent of the bulk weight is composed of seeds. The subsequent germination test indicates that 83 percent of the seeds 

germinated. Multiplying percentage purity by percentage germination yields 79 percent PLS. Illustration from Steinfeld and others (2007).
taining small seeds, or urticles, siliques, caryopses, or 

schizocarps tend to produce orthodox seeds (Hong and 

Ellis 1996, Vozzo 2002). Regardless, growers need to test 

freshly harvested orthodox seeds before long-term storage 

to develop a baseline germination percent.

After the seeds are clean, air-dry them in shallow trays for 

2 to 4 weeks before storage to reduce the moisture content. 

Stir them once a week or often enough to prevent uneven 

drying. Storing orthodox seeds requires low relative humidity, low seed-moisture content, and cool temperatures 

(figure 8.23A). A small change in seed-moisture content 

has a large effect on the storage life of seeds. With most 

orthodox species, the proper seed-moisture content for 

storage is 6 to 10 percent. Small seedlots can be stored in 

sealed jars with rubber gaskets on the lids or envelopes 

kept in a sealed, thick-walled plastic tub with an airtight lid 

(figure 8.23B). Heat-sealed plastic pouches used for food are 

effective and can be resealed as needed (figure 8.23C). An 

electronic moisture meter can be used to measure seedmoisture content and is available from several suppliers 

(figure 8.23D).

Three seed storage methods are used by small tropical 

nurseries: room temperature storage, refrigeration, and 

in some rare cases, freezing. If cooler or freezer storage is 

being used and long power outages could occur, consider 

using a backup power supply; short-term fluctuations are 

generally not a problem.

Although orthodox seeds can be stored at room temperature, they will deteriorate faster than those stored at low 

temperatures. Room temperature storage should only be 

used when seedlots are held for a short time. Seed moisture 

content during room storage needs to be at the low end of 

the range—6 to 8 percent. Seeds must be placed in airtight 

containers and stored in a room or area with low relative 

humidity. This storage method works best in the more arid 

portions of the tropics.

Collecting, Processing, and Storing Seeds 159

Figure 8.23—Orthodox seeds need to be properly dried before storage and kept in moisture-proof containers under cool conditions with 

low humidity, such as in a refrigerator at the Hawai‘i Island Seed Bank (A). Each seedlot should be labeled noting origin, date, and the viability percentage. Small lots can be stored in envelopes as long as they are kept in a moisture proof container (B) or in plastic pouches (C). 

An electronic meter measures temperature and moisture (D). Photos A, C, and D by J.B. Friday, courtesy of Jill Wagner and the Hawai‘i 

Island Seed Bank, and photo B by Tara Luna.

B

A C

D
Orthodox seeds of many tropical species can be stored 

in a refrigerator at temperatures above freezing. Seeds 

need to be placed in an airtight container and kept at 

38 to 41 °F (3 to 5 °C) in a self-defrosting refrigerator that 

maintains relative humidity between 10 to 40 percent. If 

the door is rarely opened, the humidity in a self-defrosting 

unit will maintain low relative humidity levels.

Very few tropical species will tolerate freezing—for 

those that do, dry seeds until the seed-moisture content is 

low and place them in airtight containers. When removing frozen seeds from the freezer, allow the container to 

reach room temperature before opening it. This practice 

prevents water condensation from forming on seeds.

Silica gels, available from hobby shops and florists, 

can be used to maintain low seed-moisture content with 

seeds that have been properly dried before storage. Silica 

gels have been used on short-lived native grass seeds 

placed into long-term storage to enhance longevity and 

should be tried with other short-lived seeds of native 

species (Dremann 2003). A general rule is to pour a teaspoon (about 5 ml) of silica gel into a paper envelope and 

place it with the seeds inside a tightly sealed jar for every 

2 ounces (57 g) of seeds that need to be stored. The silica 

gel will remove water vapor and ensure that seeds remain 

at the proper storage moisture. To recharge them, the 

gels can be baked in an oven (150 °F [66 °C]) for 1 hour 

or so.

Sowing Seeds After Long-term Storage

In some cases, absorbing water too quickly may damage seeds of large-seeded species that have been dried to 

low-moisture levels. Therefore, when rehydrating these 

seeds, remove them from storage and spread them evenly 

in a sealed plastic tub. Place moistened paper towels in the 

tub so that the towels do not touch the seeds directly. Water 

vapor released from the towels will be slowly absorbed by 

the seeds; after a couple of days, the seeds will be able to 

handle water uptake without injury.

References

Dawson, I.; Were, J. 1997. Collecting germplasm from trees—

some guidelines. Agroforestry Today. 9(2): 6–9.

Dremann, C. 2003. Observations on Bromus carinatus and Elymus 

glaucus seed storage and longevity. Native Plants Journal 4:61-64.

Dreesen, D. 2004. Tumbling for seed cleaning and conditioning. 

Native Plants Journal. 5: 52–54.

Dumroese, R.K.; Landis, T.D.; Wenny, D.L. 1998. Raising forest tree seedlings at home: simple methods for growing conifers 

of the Pacific Northwest from seeds. Moscow, ID: Idaho Forest, 

Wildlife and Range Experiment Station. 56 p.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

160 Tropical Nursery Manual

Seeds Are a Link Between the Evolutionary Processes of the Past and 

the Potential for Future Adaptation 

(Adapted from Flores 2002: 14–15)

“The genotype of a tree seed is the result of the evolutionary forces operating on the species for centuries. It is 

adapted to the present environmental conditions, but not to those of the future, especially when environmental 

conditions are being drastically modified by accelerated global change...The genetic combinations able to survive 

through environmental changes have intrinsic value.

Seed germination is influenced by the environmental conditions during seed development and maturation while 

on the parent tree. Day length, temperature, parental photothermic environment, light quality, and elevation are 

factors that significantly influence germination capacity. Additional factors include the inflorescence position on the 

parent tree, seed position in the fruit or infrutescence, and parent tree age during floral induction. These factors, plus 

others, explain the strong variation found in seed parameters (weight, color, water content, germinabilty) among 

seed groups and among seeds in the same group.

To most people, the concept of “seed” is deceptively simple. It is compared with a pill, isolated from the environmental effects, replicated many times, and capable of producing a plant. A seed is attached to a long and complex evolutionary and physiological history, however, and it is also conditioned to the variations of a long and complex future. 

The study of seeds has many facets and should not be limited to collection, storage, and sowing of this year’s crop.”
Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Flores, E.M. 2002. Seed biology. In: Vozzo, J.A., ed. The tropical 

tree seed manual. Agriculture Handbook 721. Washington, DC: 

U.S. Department of Agriculture, Forest Service. 13–118.

Hong, T.D.; Ellis, R.H. 1996. A protocol to determine seed storage 

behavior. International Plant Genetic Resources Institute Tech. 

Bull. No. 1. Rome, Italy: International Plant Genetic Resources 

Institute.

Kettle, C.J.; Burslem, D.F.R.P.; Ghazoul, J. 2011. An unorthodox 

approach to forest restoration. Science. 333: 36. 

Scianna, J.D. 2004. Blending dry seeds clean. Native Plants Journal. 5: 47–48.

Stein, W.I.; Danielson, R.: Shaw, N.; Wolff, S.; Gerdes, D. 1986. 

Users guide for seeds of western trees and shrubs. General Technical Report PNW-193. Corvallis, OR: U.S. Department of Agriculture, Pacific Northwest Station. 45 p.

Steinfeld, D.E.; Riley, S.A.; Wilkinson, K.M.; Landis, T.D.; Riley, 

L.E. 2007. Roadside revegetation: an integrated approach to 

establishing native plants. Technology Development Report 

FHWA-WFT/TD-07-005. Vancouver, WA: U.S. Department of 

Transportation, Federal Highway Administration. 413 p.

Thomas, D. 2003. Modifying blender blades for seed cleaning. 

Native Plants Journal. 4: 72–73.

Truscott, M. 2004. Cuisinart for cleaning elderberry (Sambucus sp. 

L. [Caprifoliaceae]). Native Plants Journal. 5: 46.

U.S. Department of the Interior, Bureau of Land Management, 

Seeds of Success[BLM/SOS]. 2011. Technical protocol for the 

collection, study, and conservation of seeds from native plant 

species for seeds of success. Native Plant Materials Development 

Program: 8–11. http://www.nps.gov/plants/sos/training/index.

htm. (October 2011).

Vozzo, J.A., ed. 2002. The tropical tree seed manual. Agriculture 

Handbook 721. Washington, DC: U.S. Department of Agriculture, Forest Service. 899 p.

Withrow-Robinson, B.; Johnson, R. 2006. Selecting native plant 

materials for restoration projects: insuring local adaptation and 

maintaining genetic diversity. Corvalis, OR: Oregon State University Extension Service.

Young, Y. 1993. Seeds of the woody plants of North America. 

Portland, OR: Dioscorides Press. 407 p.

Additional Reading

Landis, T.D.; Tinus, R.W.; Barnett, J.P. 1999. The container tree 

nursery manual: volume 6, seedling propagation. Agriculture 

Handbook 674. Washington, DC: U.S. Department of Agriculture, Forest Service. 167 p.

Schmidt, L. 2000. Guide to handling of tropical and subtropical 

forest seed. Humlebaek, Denmark: Danida Forest Seed Centre. 

511 p.

Collecting, Processing, and Storing Seeds 161
162
163

Seeds of many native species are challenging to germinate. One important thing a 

grower can do is to learn as much as possible about the life history, ecology, and habitat of the species he or she wishes to grow to understand the processes seeds from each 

target species go through in nature. Any observations will be valuable when trying 

to germinate and grow species that have little or no published information available. 

How seeds are handled, treated, and sown can affect the genetic diversity and the quality of the crop produced. Growers need to balance the desire for uniform crops and 

schedules with the need to protect the diverse characteristics within species. In this 

chapter, we discuss seed characteristics, treatments to improve or stimulate germination, and different types of sowing options for seeds.

Seed Germination and 

Sowing Options

Tara Luna, Kim M. Wilkinson, and R. Kasten Dumroese

9

Facing Page: Hand-sowing. Photo by Tara Luna.
Seed Characteristics

As discussed in Chapter 8, Collecting, Processing, and 

Storing Seeds (and shown in figure 8.8), tropical seeds can be 

divided into four categories related to their longevity and ability to be stored (Hong and Ellis 2002, Kettle and others 2011).

Viviparous: seeds that germinate before they are dispersed from the mother plant. The most common examples 

are some species of mangroves such as Avicennia and 

Rhizophora and some tropical legumes.

Recalcitrant: seeds that germinate soon after maturation and dispersal from the mother plant, and cannot be 

dried without losing viability. Most species from the wet, 

humid tropics have recalcitrant seeds because conditions 

in these environments are consistently favorable for germination and seedling establishment. Examples of common 

species with recalcitrant seeds include cacao, mango, longon, and jackfruit.

Intermediate: seeds that can germinate immediately 

but may also survive partial drying without losing viability. 

For example, papaya seeds that have been dried to 10-percent moisture content have been stored successfully under 

conditions of 50-percent relative humidity for 6 years 

without affecting viability (Vozzo 2002). Species that have 

shown intermediate storage behavior include neem, cinnamon, citrus, mahogany, and coffee.

Orthodox: seeds that can be dried without losing viability. These seeds are considered “dormant” and often 

require specific treatments to encourage germination. Dormant seeds will not germinate immediately upon maturation and dispersal from the mother plant even when ideal 

environmental conditions exist.

Before attempting to grow a plant, it is important to 

know the seed germination type because that helps determine the best seed treatments and sowing options for that 

seed. For orthodox seeds, knowing about the species helps 

you to provide the best conditions to dissipate, or “break,” 

seed dormancy and achieve good rates of germination.

Dormancy in Orthodox Seeds

Dormancy is an adaptation that ensures seeds will germinate only when environmental conditions are favorable for survival. The conditions necessary to allow seeds 

to break dormancy and germinate can be highly variable 

among species, within a species, or among seed sources 

of the same species. This degree of variability is advantageous because seeds will germinate at different times over a 

period of days, weeks, months, or even years, ensuring that 

some offspring will be exposed to favorable environmental 

conditions for survival.

Tropical species inhabiting areas with a strong wet-dry 

seasonal cycle, arid or semiarid climates, or at high elevations subjected to cold temperatures often have dormant 

seeds. The degree of dormancy in these species can vary 

among and within seed lots, between seed crop years, 

and individuals. Examples of species with dormant seeds 

include acacias such as Hawai‘i’s native Acacia koa, and 

pines including the Caribbean’s Pinus caribaea.

Dormancy may be caused by factors outside (external) 

or inside (internal) to the seeds. Some species have a combination of external and internal dormancy, a condition 

known as double dormancy. Knowing the type of seed dormancy is essential for successful propagation.

External Seed Dormancy

External seed dormancy may be physical, physicalphysiological, chemical, or mechanical (Baskin and Baskin 

1998, 2004). Seeds that have hard, thick seedcoats that physically prevent water or oxygen movement into seeds have 

physical dormancy. Physical dormancy is the most common 

seed dormancy type seen in the tropics. Species with external 

dormancy include many of the legumes (Fabaceae), mallows 

(Malvaceae), and other tropical species that are adapted 

to fire, or inhabit arid to semiarid island habitats or areas 

with pronounced wet-dry seasonal cycles. These seeds normally germinate over a period of several years. Depending 

on species and habitat, various environmental factors cause 

these seeds to become permeable over time or during a certain time of year. Seeds that require additional exposure to 

particular temperatures after they become permeable have 

physical-physiological dormancy.

The fruits that enclose the seeds cause other forms of 

external dormancy. Chemical dormancy describes fruits 

that contain high concentrations of germination inhibitors 

that prevent spontaneous germination of seeds. Mechanical 

dormancy describes tough, woody fruit walls that restrict 

seed germination and is best exemplified by the husks that 

surround coconuts (Cocos nucifera).

Internal Seed Dormancy

Internal dormancy may be morphological, physiological, 

or both (Baskin and Baskin 1998). Seeds with morphological 

dormancy have an underdeveloped embryo when dispersed 

from the mother plant. A period of after-ripening (usually 

warm and moist conditions) is needed for the embryo to fully 

mature before the seed is capable of germination. Tropical 

species that exhibit morphological dormancy are found in the 

Annonaceae, Dilleniaceae, Magnoliaceae, and Myristicaceae 

families and in many palm species. Seeds of this type may not 

germinate for several months to 1 year after sowing.

164 Tropical Nursery Manual
Physiological dormancy is found in some species 

in arid and semi-arid tropical environments. Seeds are 

permeable to water, but certain environmental conditions are necessary to modify the internal chemistry of 

the seed and thus enable germination. Usually a period 

of cold, moist conditions or holding seeds in dry storage 

overcomes physiological dormancy.

Seeds with morphological-physiological dormancy 

usually require a combination of warm and cold conditions, often over an extended period of time, before they 

are capable of germination. 

Treatments To Overcome 

Seed Dormancy and Enhance 

Germination

A variety of seed treatments have been developed in 

response to the diversity of seed types grown in nurseries. 

Before treating seeds, be sure to consult available references 

to see what treatments have been used on that species; see 

the literature cited at the end of this chapter and the Native 

Plant Network (http://www.native plantnetwork.org). If no 

information is available, check references for closely related 

species. Any personal observations made on the species in the 

habitat may also provide some clues on how to germinate the 

seeds. In general, however, the process of treating seeds follows a fairly standard progression outlined in the following 

sections. Nondormant seeds are planted immediately after 

collection and cleaning. Intermediate seeds may be stored 

for several weeks or months in suitable conditions, and then 

cleaned, fully rehydrated, and sown. For dormant (orthodox) seeds, dormancy must be overcome using one or more 

of the methods described in this chapter before seeds can be 

rehydrated, enabling germination. It is essential to determine 

which type(s) of dormancy the seed has so you can do what 

is needed to overcome dormancy (figure 9.1). The nursery 

must determine whether seeds will need to be cleaned, scarified, soaked, stimulated, stratified, and treated in other ways 

before sowing on a species-by-species basis. The following 

sections describe the seed treatment options available.

Cleaning

Seed cleaning helps prevent diseases in the nursery. Cleaning seeds of bacterial and fungal infestation is especially necessary for species that easily mold (figure 9.2). Often, molding 

can be related to the most common disease seen in nurseries, 

damping-off. Seed cleaning is especially important in humid 

climates and for species that take a long time to germinate. 

Often, without cleaning, seeds can be lost to pathogens before 

they are planted in the nursery.

One of the best cleaning methods is to simply soak seeds 

in a stream of running water for 24 to 48 hours. The running water flushes bacterial and fungal spores from the seeds 

(James and Genz 1981). This treatment can also be used to 

satisfy the soaking requirement described in the next section.

Seeds can also be cleaned with several chemicals, some 

of which also act to stimulate germination. Bleach (5.25 

percent sodium hypochlorite) is the most common chemical used. Depending on the species, bleach cleaning solutions range between one part bleach in eight parts water to 

two parts bleach in three parts water. With most species, 

Seed Germination and Sowing Options 165

Figure 9.1—Key to dormancy types. Knowing the type of seed 

dormancy is essential to successful seed propagation. Illustration 

by Jim Marin.

Figure 9.2—Seeds that are not cleaned before treatment or 

sowing can easily mold or be susceptible to serious pathogens 

such as damping-off disease. Photo by Thomas D. Landis.
treatment duration is 10 minutes or less. Species with very 

thin seed coats should not be cleaned with bleach. Hydrogen peroxide can be an effective cleanser and can sometimes enhance germination (Narimanov 2000). The usual 

treatment is one part peroxide in three parts water. Tropical 

species that benefit from hydrogen peroxide rinses include 

Albizia species and camphor tree seeds (Vozzo 2003).

Scarification

Seeds with external dormancy require scarification. 

Scarification is any method of disrupting an impermeable 

seed coat so that water and oxygen can enter the seeds. 

In nature, hard seed coats are cracked or softened by fire, 

extreme temperatures, digestive acids in the stomachs of 

animals, or by the abrasion of blowing sand. After the seed 

coat has been disrupted, oxygen and water pass into the 

seeds and germination can proceed.

Seeds can be scarified many ways. How well the method 

works depends on the species and the thickness of the seed 

coats. Whichever method is chosen, it is very important not 

to damage the endosperm, cotyledons, or embryo during 

the treatment. Taking time to learn seed anatomy of the 

species is helpful. Trying several methods and recording 

the results will help determine the best method for that species and seed source.

Mechanical Scarification

Mechanical scarification includes filing or nicking seeds 

by hand and is most often used on large-seeded species such 

as Acacia, Cassia, and Sesbania (figure 9.3). Be sure to scarify on the side of the seed opposite the embryo. It is often 

done one seed at a time with a nail clipper. This method is 

time consuming and requires precision to adequately scarify 

the seed coat without damaging the internal portions of the 

seed. Sandpaper can be used on smaller seeded species such 

as sedges; placing seeds into a shallow wooden box and then 

rubbing them under a block of wood covered in sandpaper is 

the simplest technique. Often, however, the degree of scarification achieved with sandpaper can be variable.

Hobby-size rock tumblers can be used to process large 

batches of seed more quickly than manual mechanical 

scarification (figure 9.4). Dry tumbling involves placing 

seeds, a coarse carborundum grit (sold by rock tumbler 

dealers), and pea gravel in the tumbler and tumbling for 

several hours or several days. Wet tumbling includes the 

addition of water to the grit and pea gravel. A benefit of wet 

tumbling is that seeds are soaked in well-aerated water and 

chemical inhibitors may be leached from the seed.

Heat Scarification

Many species, especially those from fire-adapted ecosystems, respond to germination cues from heat. Using 

either wet or dry heat to scarify the seeds can simulate this 

response. Using wet heat is an effective method for many 

small-seeded species because it provides a rapid, uniform 

treatment that can be assessed within a few hours. Wet166 Tropical Nursery Manual

Figure 9.3—Mechanical scarification works well with large, 

easy-to-handle seeds. Great care is needed, however, so that the 

embryo or cotyledons are not damaged. Koa seed hand scarification (A), scarified lotus seeds (B). Photo A by Craig R. Elevitch, 

and photo B by Cardno JFNew Native Plant Nursery.

Figure 9.4—Hobby-size rock tumblers can be used to scarify seeds 

and avoid seed destruction that can occur with sulfuric acid or heat 

scarification. Photo by Tara Luna.

B

A
heat treatments are effective for many tropical species 

including Acacia, Cassia, Senna, Sesbania, and Tamarindus (Vozzo 2002). Because the thickness of the seed 

coat may vary among sources, it is wise to dissect a few 

seeds and examine the thickness of their seed coats to 

help determine treatment duration. Seeds are added to 

boiling water for 5 to 10 seconds and then immediately 

transferred to a vat of cold water so that they cool quickly 

to prevent embryo damage. Seeds imbibe the cool water 

for 1 day and are then ready for sowing or for stratification (figure 9.5). Some species cannot tolerate excessively 

high temperatures, so you may want to heat the water to 

only 158 °F (70 °C) and monitor your results.

Dry heat is most commonly used on fire-adapted species. Seeds are placed in an oven at temperatures ranging 

from 175 to 250 °F (80 to 120 °C) from a few minutes to 

1 hour, depending on the species. The seed coat cracks 

open in response to the heat. To avoid damaging seeds, this 

treatment needs to be monitored closely.

Chemical Scarification

Sulfuric acid is most commonly used on species with very 

thick seedcoats and with stony endocarps that surround the 

embryo (figure 9.6). It has been used on some species of Acacia, Albizia, Cassia, Leucaena, Parkinsonia, and Terminalia 

(Vozzo 2002). Treatment length varies with the species and 

often among seed sources, and it must be carefully monitored 

because seeds can be destroyed if the treatment is too long. 

A simple way to monitor the process is by removing seeds at 

regular intervals and cutting them with a sharp knife. When 

the seeds are still firm but can be cut fairly easily, the treatment is probably sufficient. Another way is to run a pilot test 

on a subsample of seeds. Again, remove some seeds periodically and evaluate how well they germinate. After the best 

duration is known, the entire seedlot can be treated. Sulfuric 

acid is very dangerous to handle and requires special equipment, personal protective gear, and proper disposal after use. 

It should never be poured down sink drains. If acid is being 

diluted in water, the acid must be added to the water, never 

add water to acid—when water is added to acid, heat will 

be released, risking an explosion and other dangers. Some 

species have thick seed coats but can easily be damaged by 

sulfuric acid. Instead, citric acid or sodium or calcium hypochlorite baths with longer treatment durations may be used.

The safe use of sulfuric acid requires the following procedures:

• Treat seeds that are dry and at room temperature.

• Require workers to wear safety equipment, including face shield, goggles, thick rubber gloves, and full 

protective clothing.

• If diluting, add acid to water, never water to acid.

• Immerse seeds in an acid-resistant container, such as 

a glass, for the duration required.

• Stir seeds carefully in the acid bath; a glass rod 

works well.

• Immerse the container with seeds and acid in an 

ice bath to keep temperatures at a safe level for the 

embryos (this temperature depends on the species; 

many do not need this step).

• Remove seeds from the acid by slowly pouring the 

seed-acid solution into a larger volume of cool water, 

ideally one in which new, fresh water is continually 

being added.

• Stir seeds during water rinsing to ensure all surfaces 

are thoroughly rinsed clean.

Seed Germination and Sowing Options 167

Figure 9.6—Seeds that have been treated with sulfuric acid. Photo 

by Nancy Shaw.

Figure 9.5—Seeds that have been scarified by hot water are 

visibly larger than untreated seeds because the seed coat has been 

breached and seeds can then absorb water increasing their size. 

Photo by Greg Morgenson.
Soaking

After cleaning and scarification, seeds must have exposure to water and oxygen before germination can occur. The 

standard procedure is to soak seeds in water for 1 to several 

days until they are fully hydrated (figure 9.7). Hydration 

can be checked by taking a sample, allowing it to dry until 

the seed coat is still wet but dull, not glossy, and weighing 

it. When the weight no longer increases substantially with 

additional soaking time, the seeds have absorbed sufficient 

water. Scarified seeds will be more obvious; the seeds will 

enlarge drastically during the soak. Seeds that only had physical dormancy can be immediately planted. As mentioned 

previously, running water rinses are effective seed cleaning 

treatments that reduce the need for fungicides in nurseries 

(Dumroese and others 1990). Running water soaks also help 

to remove any chemical inhibitors present on or within the 

seeds. An aquarium pump can be used to agitate the seeds to 

improve the cleaning effect and keep the water well aerated. 

If seeds are not soaked with running water, change the water 

often (at least a couple of times each day).

Germination Stimulators

Several chemicals are known to increase seed germination. These chemicals are usually applied after seeds are 

fully hydrated. In general, only seeds with internal dormancy 

receive this treatment. Germination stimulators include gibberellic acid, ethylene, smoke, and potassium hydroxide.

Gibberellic Acid

Gibberellic acid is the most important plant hormone for 

the regulation of internal seed dormancy and is often used 

on seeds with complex internal dormancy and with those 

species having underdeveloped embryos. In some cases, 

it has been used to substitute for a warm, moist treatment and to hasten embryo after-ripening. Sandalwood 

is a species that has been successfully germinated using 

gibberellic acid. Gibberellic acid can be purchased from 

horticultural suppliers. Preferred concentrations vary, 

but most nurseries use 500 to 1,000 parts per million 

(ppm). High concentrations can cause seeds to germinate, but the resulting seedlings may be of poor quality. 

Therefore, it is best to experiment with low concentrations first. The following are some guidelines for treating 

seeds with gibberellic acid:

• Gibberellic acid takes a long time to dissolve. It may 

need constant stirring or you may want to prepare it 

the day before use.

• Store unused solution away from direct sunlight.

• Cut unbleached coffee filters into squares and fold 

them diagonally.

• Place gibberellic acid solution evenly into an ice cube 

tray.

• Place each folded coffee filter containing the seeds 

into the wells of the tray so that it wicks up the solution.

• After 24 hours, remove and either sow directly or 

place seeds into fresh coffee filters moistened with 

distilled water for stratification.

Ethylene

This gas occurs naturally in plants and is known to stimulate the germination of some species. Ethylene gas is released 

from ethephon, a commercially available product. Ethephon, 

used either alone or in combination with gibberellic acid, has 

enhanced the germination in doum palm (Mousa and others 

1998) and may be used for other species inhabiting arid to 

semi-arid tropical and saline environments. It may inhibit 

germination in other species, so consult the literature and 

experiment before using operationally.

Smoke Treatments

Smoke stimulates germination in many fire-adapted 

species; for example, species from the California chaparral, 

longleaf pine communities in Florida, or species from firedependent ecosystems in Australia, South Africa, parts of 

South America, and the Mediterranean. Smoke especially 

stimulates seeds that have thin, permeable seed coats that 

allow entry of smoke into the seeds (Keeley and Fotheringham 1998). Seeds can be treated with smoke fumigation, a 

method in which smoke is piped into a specially constructed 

smoke tent containing seeds sown in trays (figure 9.8A), 

168 Tropical Nursery Manual

Figure 9.7—Many species of dormant seeds benefit from 1 to several days of water soaking before sowing to fully imbibe seeds and 

remove any chemical inhibitors within the seeds or on the seed coats. 

Photo by Brian F. Daley.
or with smoke water. Smoke water is an aqueous solution 

of smoke extract made by burning vegetation and piping 

the smoke through distilled water or allowing the smoke 

to infuse into a container of water. Seeds are then soaked 

in the treated water (figure 9.8B). Conversely, growers can 

experiment with commercially available smoke products 

such as liquid smoke or smoke-infused paper discs, or by 

adding ash to growing media.

Many variables, such as the material used for combustion, the combustion temperature, and the duration of 

exposure, will need to be determined on a species-by-species basis. Experiments performed by Keeley and Fotheringham (1998) found that the length of exposure to smoke 

was very important in some species; a 3-minute difference 

in exposure resulted in seed mortality. Some fire species 

did not germinate under heat or smoke treatments alone. 

With some species, seed burial for 1 year or stratification 

was required in addition to smoke exposure. All these 

factors can have an effect on germination and should be 

considered when determining whether to use smoke treatments. Success with this novel treatment will require trials, 

so good record keeping is critical.

Potassium Hydroxide Rinses

Potassium hydroxide has been used to stimulate germination in several native plant species. Optimum concentration varies from 5.3 to 7.6 Molar for 1 to 10 minutes 

depending on the species; longer soaks at higher concentrations were found to be detrimental (Gao and others 1998).

Other Stimulants

Potassium nitrate, thiourea, and kinetin have been used 

to stimulate germination in seeds, although the use of these 

compounds with tropical native plants is lacking. You may 

choose to experiment with these compounds on a limited 

basis with difficult-to-germinate seeds.

Temperature and Moisture Treatments

Many seeds with internal dormancy require a moist 

period at certain temperatures similar to what occurs in 

the natural habitat before they germinate and grow. Historically, stratification was a temperate zone practice of 

alternating layers of moist soil and seeds in barrels and 

allowing these “strata” to be exposed to winter temperatures. Nowadays, stratification is often used more generically to describe the combined use of moisture and any 

temperature to overcome seed dormancy. We use the term 

“stratification” to refer to only cold, moist treatments (rare 

to use in the tropics, but we cover it anyway for nurseries 

growing highland species). We use the term “warm, moist 

treatment” instead of “warm, moist stratification.”

Some native species with double internal seed dormancy 

require a combination of a warm, moist treatment for a 

period of time followed by stratification. Some species or 

seedlots may require only a few days or weeks of stratification, while others may require several months. As a general 

rule, it is best to use the maximum recommended treatment. 

Also keep in mind that what works well at one nursery may 

not necessarily work well at another nursery because of differences in seed source, handling, processing, cleaning, and 

storage. An advantage to stratifying seeds of some species is 

that it can speed up germination and make it more uniform, 

which is desirable in a container nursery.

Seed Germination and Sowing Options 169

Figure 9.8—Smoke treatments have been used to overcome seed 

dormancy and enhance germination rates for many native species 

inhabiting fire-dependent ecosystems. A smoke tent for treating 

seeds (A). Smoke water-treated seeds of angelica (B). Photo A by 

Kingsley Dixon, and photo B by Tara Luna.

B

A
Figure 9.9—Small seeds requiring only a few weeks of stratification can be stratified by moistening paper towels and holding by corner to 

let excess water drain away (A) or placing seeds onto moistened towels inserted into an unopened plastic zippered bag (B). Illustrations from 

Dumroese and others (1998).

Warm, Moist Treatments

Warm, moist treatment enhances after-ripening of 

seeds with underdeveloped embryos. Warm, moist treated 

seeds are kept at temperatures of 72 to 86 °F (22 to 30 °C) 

for a period of time, usually in moist peat moss, sawdust, or 

other substrate. Although warm, moist treatments are not 

commonly used on tropical species, it can be considered for 

seeds with morphological or physiological seed dormancy.

Stratification

Stratification (cold, moist treatment) is used on seeds with 

internal dormancy from temperate areas, or high-elevation 

habitats in tropical regions. Some subtropical species may also 

benefit from a period of cool, moist stratification. In climates 

with four seasons, seeds sown in flats or containers in late 

summer or autumn and left outdoors during winter undergo 

“natural” stratification. This technique may be preferred if the 

species has double dormancy (requires both a warm, moist 

treatment or stratification), requires a very long stratification 

or requires low temperatures or fluctuating temperatures for 

a long period of time. Conversely, “artificial” stratification 

involves placing seeds under refrigeration at 34 to 38 °F (1 to 

3 °C) for a period of time. Artificial stratification has several 

advantages: (1) it allows for a routine check of seeds to ensure 

they are moist and not moldy, (2) a large number of seeds can 

be stratified in a small space, and (3) seeds or seedlots that 

begin to germinate can be removed from the treatment and 

planted in the nursery as they become available. Artificial 

stratification is preferred over natural stratification unless the 

natural treatment provides higher rates of germination.

For artificial stratification of small seedlots and small 

seeds, seeds can be placed between sheets of moistened 

paper towels and inserted in an opened plastic bag or sown 

on a medium in flats with drainage holes. Paper towels 

need to be moist but not waterlogged, and seeds need to be 

evenly spread across the moist paper towel to help prevent 

molding (figure 9.9).

Another technique is “naked” stratification. Most conifer 

seeds, for example, are stratified in this manner (figure 9.10). 

Seeds are placed in mesh bags and then soaked in running 

water as described previously. After the seeds are hydrated, 

the bag is pulled from the soak, allowed to drip dry for 30 to 

90 seconds, and then suspended in a plastic bag. Make sure 

the seeds are not in contact with standing water in the bag 

and hang the bags in the refrigerator. If naked seeds need a 

warm, moist treatment before stratification, it is easiest to 

first spread the seeds onto moistened paper towels enclosed 

in large plastic bags. After the warm treatment, the seeds can 

be returned to the mesh bags for stratification. One other 

hint: if a particular species or seedlot has a tendency to begin 

germinating during stratification, surface-dry the seed coats 

(seeds should be moist and dull, not shiny), and then put the 

seeds into the bag for refrigeration. The seeds need to still 

have enough moisture for chemical processes that dissipate dormancy to occur but not enough moisture to allow 

for germination.

170 Tropical Nursery Manual

A B
Figure 9.10—Naked stratification: After soaking seeds in a mesh bag, allow the bag to drip dry for 1 minute and then suspend the mesh 

bag in a plastic bag (A). Hang the bag in the refrigerator. Make sure the seeds are not in constant contact with standing water in the plastic 

bag (B). Illustrations from Dumroese and others (1998).

Many wetland and aquatic species can be treated with 

naked stratification in water. In general, these species can 

be easily stratified in Ziploc®-type bags filled with water. 

Insert a soda straw into the bag, ensuring that the end is 

sticking out of the bag, to allow some oxygen to reach the 

seeds. Then, seal the rest of the bag securely. Place under 

refrigeration if in need of a cold, moist stratification period.

Environmental Factors 

Influencing Germination

Four environmental factors affect germination: light, 

water, oxygen, and temperature. All plants have specific 

germination requirements based on ecological adaptations 

and the environmental cues that trigger germination for 

that species.

Light

Light quality and duration can influence germination. 

In nature, seeds of tropical pioneer species require high 

light levels associated with a canopy gap for germination and establishment, whereas shade tolerant species 

generally can germinate in very poor light or deep shade. 

Many small-seeded, tropical native species fall into this 

category. Thus, pioneer species, such as ‘ōhi‘a in Hawai‘i, 

with very small dust-like seeds (figure 9.11), require light 

for germination and fail to germinate even if they are buried only 2 mm deep (Drake 1993). Therefore, these seeds 

need to be sown on the surface of the medium so they are 

exposed to light during germination. Other species are 

conditioned to germinate only if they are buried in the 

soil. Species requiring darkness to germinate are those 

that germinate readily under the deep shade of a closed 

forest canopy. Tropical trees and shrubs with medium to 

larger sized seeds often require darkness for maximum 

germination, but shade tolerant vines and herbaceous 

plants may have smaller seeds. Other species requiring 

darkness to germinate include some of the species that 

colonize sand dunes along coastlines.

Seed Germination and Sowing Options 171

Figure 9.11—In general, small-seeded pioneer species such as 

‘ōhi‘a require light to germinate and must not be buried. Photo 

by Tara Luna.

A B
Water and Oxygen

Water is also important for germination. Overwatering seeds during germination results in reduced levels of 

oxygen in the medium and promotes tissue breakdown 

and disease whereas underwatering delays or prevents 

germination. Therefore, seeds need to be kept evenly 

moist during germination. Although oxygen is needed 

for respiratory processes in germinating seeds, some 

aquatic species may require low oxygen levels for germination. For example, tropical floodplain forest or wetland 

species naturally germinate during periods of high water 

inundation and respond positively to low water oxygen 

levels (Kurbitzky and Ziburski 1994, Vozzo 2002).

Temperature

Temperature influences seed germination rate and 

percentage. Some germination patterns in response to 

temperature include seeds that require cool temperatures, tolerate cool temperatures, require warm temperatures, and (or) require alternating temperatures (Hartman and others 1997, Vozzo 2002). Species requiring 

cool temperatures generally germinate below 77 °F (25 

°C), which coincides with high elevations in tropical or 

subtropical regions. Species that tolerate cool temperatures will germinate over a wide range of temperatures 

from 41 to 86 °F (5 to 30 °C). Many species will not 

germinate under excessively high temperatures. Most 

tropical species require warm temperatures and will only 

germinate if temperatures are above 70 °F (21 °C). In 

addition, some species germinate better when exposed to 

alternating temperatures. Alternating temperatures are 

particularly important with dormant, freshly harvested 

seeds. Many tropical tree species germinate to their highest percentages at alternating temperatures of 86/68 °F 

(30/20 °C) or are provided with at least 10 °F (5.6 °C) 

difference between the day and night time temperature. 

Some difficult-to-germinate tropical species may require 

even greater temperature fluctuations.

Seed Sowing Methods

Several sowing techniques have been used for native 

plants (table 9.1) and are described in the following sections. The process of sowing seeds for nursery production 

will vary with the species, type of seed, seed quality, and 

nursery environment.

Direct Sowing

Direct sowing is fast, easy, and economical because 

it minimizes seed handling and labor. It can be mechanized when done on a large scale. For direct sowing to be 

efficient, the seeds need to be easy to handle, abundant 

in supply, have simple dormancy treatments, and have a 

known high-germination rate (figure 9.12). If the direct 

sowing will be mechanized, seeds must also be uniform in 

size and shape.

The success of direct seeding depends on the accuracy 

of seed germination information. Growers must realize 

that actual seedling emergence may be different from 

the results of laboratory germination tests that are conducted under ideal environmental conditions. Nursery 

managers must adjust for this discrepancy based on their 

own operational experience. Growers should conduct a 

small germination test of each seedlot to determine the 

172 Tropical Nursery Manual

Figure 9.12—Direct sowing works well 

for seeds that have little or no dormancy 

(or have been treated to overcome dormancy), are easy to handle and are in 

abundant supply (A). Simple tools like a 

film canister (B) or a folded envelope (C) 

can be used to accurately sow small seeds. 

Photo A by Douglass F. Jacobs, and photos B and C by Dawn Thomas.

B

A C
percentage of germination for each seedlot. Those percentages can then be used to determine the number of 

seeds to direct sow (see table 9.2). Follow these steps for 

successful direct sowing:

• Determine how many seeds must germinate to obtain 

the production target.

• Determine if seeds can be single-sown or will require 

multiple seeds to reach the production target (see 

following sections).

• Cleanse and treat seeds as necessary to break dormancy.

• Sow seeds, ideally centering the seeds in each container. Some seeds require a specific orientation for 

optimal growth and development; if so, make sure 

seeds are sown in the correct orientation.

• Depending on the light requirements of the species, 

cover seeds with the correct amount of mulch.

• Gently water the seeds with a fine watering head to 

press them into the growing media.

Multiple-Seed Sowing and Thinning

Sowing more than one seed into each container 

with the expectation that at least one will germinate is 

the most common direct-sowing practice. The number 

of seeds to sow can be calculated based on the seeds’ 

Seed Germination and Sowing Options 173

Table 9.1—Methods for sowing seeds. Adapted from Landis and others (1999). 

Propagation method

Good method for 

seeds with the following 

characteristics

Advantages Disadvantages

Direct Sowing:

Seeds are sown into 

containers

• Have a known highpercentage germination

• Are inexpensive

• Are in abundant supply

• Have uniform, smooth shapes

• Fast and easy

• Economical

• Minimizes seed handling

• Seeds are all sown at once

• Less efficient use of space, seeds, 

and/or growing medium

• Causes of poor germination are 

difficult to track

• May require thinning 

and/or consolidation and associated labor costs

• Not good for large or 

irregularly shaped seeds

Planting Germinants:

Seeds sprouting or 

germinating in trays or 

bags are sown into containers 

while roots are just beginning 

to emerge

• Are of unknown viability

• Are valuable or rare

• Have unknown germination 

requirements

• Germinate during an extended 

period of time or during stratification

• Efficient use of seeds

• Efficient use of nursery space

• Can adjust for unknown 

seed quality or performance

• Labor intensive

• May result in nonuniform crop 

development

• Root deformation possible

• Requires frequent, skilled 

monitoring

Transplanting emergents:

Seeds are sown into flats or 

seedbeds for germination; once 

germinated and leaves appear, 

seedlings are transplanted to 

containers

• Are being tested but will not be 

transplanted to produce a crop

• Do not respond well to other 

sowing methods

• Have long or unknown 

dormancy

• Good for trials to observe seed 

performance

• Useful with fibrous 

rooted species

• Efficient use of seeds

• Efficient use of nursery space

• Can adjust for unknown seed 

quality or performance

• Not recommended for 

woody and/or taprooted 

species because of problems 

with transplant shock and 

or root deformation

• Requires skilled labor

Miniplug transplants:

Seeds are sown directly 

into small containers. 

After germination, they are 

transplanted into larger 

containers

• Are of unknown quality

• Are valuable or rare

• Have unknown germination 

requirements

• Have very tiny seeds

• Will be transplanted into 

large containers

• Efficient use of space

• Uniform crop development

• Low risk of transplant injury

• Requires two sets of 

containers

• Timing is critical

• Transplanting by hand 

is labor intensive
expected germination percentage. Two to five seeds are 

typically sown per container. As a general rule, seeds 

with less than a 50-percent germination are not recommended for direct sowing because the high density of 

nonviable seeds in the container may cause disease problems, more containers will need to be thinned, and many 

plants will be wasted (figure 9.13). Table 9.2 provides 

general recommendations for the number of seeds to 

sow per container based on the germination percentage. 

At some point, adding more seeds per container does not 

really increase the number of containers with plants (table 

9.3) but does drastically increase the number of containers with too many plants and the amount of seed wasted. 

Sometimes it may be better to single-sow a few containers 

than thin extra seedlings from many containers. For example, sowing a single seed per container of a seedlot with 

85-percent germination yields 15 percent empty containers 

whereas sowing two seeds per container yields only 2 percent empty containers, but sowing the extra seed requires 

174 Tropical Nursery Manual

Seeds sown 

per container Empty containers (%) Containers with at

least one seedling (%)

Seed

sown 

Seedlings

produced

1 35 65 1,000 650

2 12 88 2,000 880

3 4 96 3,000 960

4 1 99 4,000 990

5 0 100 5,000 1,000

Seed germination percentage Seeds to sow per container Percentage of containers with 

at least one seedling

90 + 1 to 2 90 to 100

80 to 89 2 96 to 99

70 to 79 2 91 to 96

60 to 69 3 94 to 97

50 to 59 4 94 to 97

40 to 49 5 92 to 97

Table 9.3—A sowing example for a seedlot of Acacia koa having a 65-percent germination rate. Assuming 1,000 seedlings are 

desired, notice that adding more than three seeds per container really does not improve the number of containers with seedlings and 

wastes many seeds. Adapted from Dumroese and others (1998).

Table 9.2—For a given seed germination, increasing the number of seeds sown per container increases the number of filled 

containers. In general, a target of 90- to 95-percent filled containers is reasonable. Adapted from Dumroese and others (1998).

Figure 9.13—Calculating and testing germination rates help 

reduce costs and problems associated with thinning. Photo by 

Thomas D. Landis.
thinning 72 percent of the containers. The nursery manager 

may have been better off, in terms of seed use efficiency and 

labor, to have simply oversown 10 percent more containers 

rather than pay for the labor to thin. Therefore, the amount 

of seeds to sow per container is a function of germination, 

seed availability, nursery space, thinning costs, and so on.

When more than one seedling germinates in the same 

container, seedlings compete for light, water, and nutrients. This competition results in lower initial growth rates 

and requires that seedlings be thinned (clipped, culled, or 

removed from the container). For this reason, thinning 

should be done as soon as possible after seedlings emerge. 

Thinning is a labor-intensive practice and it can damage 

remaining seedlings if done improperly. Train workers to 

thin plants carefully and to follow these guidelines:

• Thin germinants as soon as possible; the more developed 

the root system becomes, the more difficult it is to thin.

• Retain the strongest seedling closest to the center of 

the container. Thinning is an opportunity for selecting the healthiest seedling while removing inferior 

plants.

• Pull or cut extra plants. For species with a long, slender taproot at germination (such as pine seedlings), 

extra seedlings can be easily pulled before they develop 

secondary roots. For species with vigorous, fibrous root 

systems, cutting extra plants at the stem with sharp 

scissors or nipping them with fingernails may be better.

• Discard culled plants into compost or waste.

• Check the remaining seedling and correct any disruptions caused by the thinning process. (For example, if 

thinning disrupted the mulch, adjust it so the seedling 

has the best environment possible.)

Single-Seed Sowing

Sometimes, particularly when seeds are scarce or costly or 

are expected to have close to 100-percent germination, single seeds can be directly sown into containers. This practice 

ensures that every seed has the potential to become a plant 

and no thinning will be necessary. If a particular number of 

plants are required, then extra containers are planted, often 

referred to as “oversowing,” to make up for any empty cells. 

The number of extra containers to sow can be calculated 

based on the percentage of germination. If a seedlot has only a 

78-percent germination, for 100 plants, you must sow at least 

28 extra containers (100 desired seedlings/0.78 success rate = 

128 containers required). The number of oversown containers may need to be increased to account for seedling losses 

during the growing cycle.

Oversowing works best if the nursery has extra space and 

is using containers with individual, exchangeable cells because 

containers with live plants can be consolidated and the extra 

containers can be removed (see Chapter 7, Containers). Single 

sowing is efficient because no seeds are wasted and plants that 

do emerge are not subjected to competition or the stresses of 

thinning as they are with the multiple-sowing technique. Oversowing wastes potting materials and bench space, however, 

and consolidating the empty containers is labor-intensive.

Sowing Germinants

Germinant sowing (“sowing sprouts”) is the practice 

of sowing seeds that are germinating (or sprouting) into 

the container when their young root emerges (figure 

9.14). When done properly, germinant sowing ensures 

that one viable seed is placed in each container, thereby 

making efficient use of space and seeds. The resulting 

seedlings are often larger because they can begin to grow 

immediately without competition. This technique can be 

labor-intensive but results in minimal waste of materials 

and space. Sowing germinants works best for seeds that:

• Are from a rare or valuable seedlot.

• Have a low or unknown germination percentage.

• Are large or irregularly shaped.

• Germinate in stratification.

• Have deep dormancy and germinate over a long 

period of time.

• Rapidly produce a long root after germination (such 

as many desert and semidesert species).

Germinant sowing is a relatively simple process. Seeds 

are treated as necessary, then germinated in trays or bags. 

Seed Germination and Sowing Options 175

Figure 9.14—Germinants must be sown as soon as the radicle 

emerges from the seed coat. Photo by Tara Luna.
Seeds may be spread out between layers of moist paper towels or moist cardboard. Larger seeds are sometimes placed 

in plastic bags filled with a moist medium such as Sphagnum peat moss. Seeds are closely spaced, but far enough 

apart so that mold does not spread if it forms. Seeds are 

checked every few days. After seeds begin to germinate, 

they must be checked daily. Germinated seeds are removed 

daily and planted directly into growing medium in their 

containers. Larger seeds can be planted by hand; smaller 

seeds are often sown using tweezers.

Timing and root orientation are critical when sowing 

germinants. Seeds need to be sown into containers as soon 

as the root emerges. The embryonic root, often called a 

“radicle,” needs to be short, ideally no longer than 0.4 in (1 

cm). If the radicle becomes too long, it may be difficult to 

plant without causing root deformation (figure 9.15). Some 

growers like to prune the radicle of taprooted species before 

planting to ensure a more fibrous root system. No more 

than the very tip (up to 0.1 in [3 mm]) is trimmed with 

clean scissors. The germinating seed is carefully placed in 

the container with the radicle extending downward. After 

the seeds are properly planted, the medium needs to be 

firmed around the root and the seed covered with mulch.

An advantage of planting germinants is that the germination process is more visible to the growers than when 

seeds are direct sown. Germination timing can be better 

monitored and the causes of germination problems are easier to track, but because seeds in trays or bags are very close 

together, a mold or pathogen can contaminate all the seeds 

if not properly monitored. Labor is required to routinely 

check for germination, skill is required to achieve proper 

planting orientation of the seeds, and planting must be 

done in a timely fashion. Because germinants may emerge 

over several weeks or longer for some species, crop development will be more variable and require special cultural 

treatments.

Transplanting Emergents

Transplanting emergents (“pricking out”) is a practice 

for germinating seeds in a small area. Seeds are hand-sown 

in shallow trays that are usually filled with about 2 in (5 cm) 

of peat moss-vermiculite (or similar) growing medium (figure 9.16). Soon after the seeds germinate, they are “pricked 

out” of the tray and transplanted into a container. This 

technique is not recommended for woody plants and other 

taprooted species, because root problems often result.

Transplanting emergents works best when—

• Species have a fibrous root system that recovers well 

from transplanting (herbaceous forbs without a taproot, grasses, sedges, and rushes).

176 Tropical Nursery Manual

Figure 9.15—When planting germinants, seeds must be sown 

as soon as the radicle is visible and must be oriented correctly 

when planted. Incorrect orientation leads to severe root deformation in woody species (A, B). Photo A by Thomas D. Landis, and 

photo B by R. Kasten Dumroese.

Figure 9.16—For transplanting emergents, seeds are handsown in trays that are usually filled with about 2 in (5 cm) of 

peat moss-vermiculite growing medium. To prevent root deformities, it is important the trays be deep enough that the roots 

will not touch the bottom before the emergents can be transplanted. Photo by Thomas D. Landis.

B

A
• Tests or trials are being used to observe seed treatments, germination timing or percentage, early 

growth rate or other early developmental issues.

• Seeds are too small or fragile to be sown by any other 

method.

• Seeds have very complex dormancy or germinate over 

an extended period of time.

• Limited nursery growing space makes direct seeding

uneconomical.

• Timing to transplant emergents is scheduled 

promptly.

Some key disadvantages include the following—

• Disease potential is high in densely planted trays.

• Root orientation and timing is critical; root malformations and other problems can result if neglected.

• Transplanting is skill and labor intensive. 

Training, care, good timing, and proper technique is 

required to prepare seedling trays, sow the seeds, and to 

transplant emergents properly. As with many other nursery 

functions, some trial and error occurs in finding the medium 

mixture and tray depth that works best for each species. 

Larger seeds are scattered by hand over the surface of the 

moistened medium, or place in indentations in the medium. 

Smaller seeds can be sown with a salt shaker with enlarged 

holes. Sown seeds are then covered with a light application 

of fine-textured mulch or medium, irrigated, and placed in a 

favorable environment for germination. Although the exact 

size or age to transplant the germinating seedlings varies 

by species, it is usually done at the primary leaf stage (after 

cotyledons emerge) and well before root systems reach the 

bottom of the seed tray (figure 9.17).

Emergents are carefully removed from the tray, usually 

by gently loosening the medium around them (figure 9.18A). 

A small hole is made in the medium of the container and the 

germinant is carefully transplanted, ensuring proper root 

orientation (figure 9.18B). Some species benefit from 

root pruning before transplanting. The potting medium 

is then firmed around the root and stem (figure 9.18C).

Seed Germination and Sowing Options 177

Figure 9.17—Root and shoot development at various stages 

after germination of the Caribbean coastal shrub, Chrysobalanus icaco. The large-seeded species is easily transplanted at 

these stages. Photo by Brian F. Daley.

Figure 9.18—Transplanting emergents works well for fibrous-rooted shrubs, forbs, and grasses. Great care must be taken to lift 

the emergent from the pricking out tray without damaging the roots (A) and to carefully and properly transplant it into the new 

container filled with moistened growing media (B, C). Photos by Tara Luna.

A B C
When timed incorrectly or done improperly, especially 

on taprooted woody species, transplanting emergents can 

produce a “J-root” or kink in the seedling stem or root 

(figure 9.19). These malformations can cause mechanical 

weakness, poor growth in the nursery and later in the field, 

and mortality after outplanting. Therefore, unless no other 

sowing method works, transplanting emergents of woody 

plants is discouraged.

Transplanting Plugs

Small-volume containers, such as miniplugs (figure 

9.20) or expanded peat pellets, in which seeds are direct 

sown (see Chapter 7, Containers) can be transplanted 

into a larger container after the seedlings are well established. Transplanting small plugs has a number of benefits. The small plug container preserves healthy root 

form because damage to roots during transplanting is 

eliminated. Planting small plugs also makes efficient use 

of growing space. Large numbers of small plugs can be 

started in a very small area and managed intensively during germination and early growth.

Plants in miniplug containers must have a firm enough 

root plug to hold the plug together and withstand the 

transplanting process, but they must not have so many 

roots that they are rootbound or the roots may become 

deformed after transplanting. If peat pellets are used, too 

few roots are not a problem because the entire pellet can 

be transplanted. A hole large enough to accept the plug 

is made in the medium of the larger container, and the 

small plug-grown seedling is carefully inserted. Planters 

need to ensure that the roots go straight down and are 

not deformed during transplanting. The medium is gently firmed around the root system, mulch is applied, and 

the plant is watered.

Transplanting small plugs is labor intensive and 

requires skill. In some arid, windy areas, small plugs are 

not practical because they dry out too quickly between 

waterings. Before investing in small plugs on a large scale, 

a small trial is advised.

Seed Coverings (Mulch)

Regardless of the seed sowing method, a seed cover 

or “mulch” is recommended to create an optimal environment for germinating seeds. The only exception is for 

species that require light to germinate. Mulch is usually a 

light-colored, nonorganic material spread thinly over the 

seeds. Examples of mulches include granite grit (such as 

poultry grit) (figures 9.21A, 9.21B), pumice, perlite (figure 9.21C), coarse sand, or vermiculite (figure 9.21D). 

When properly applied, mulches—

• Create an ideal “moist but not saturated” environment around germinating seeds by making a break 

in the texture of the potting medium (water will not 

move from the medium into the mulch).

178 Tropical Nursery Manual

Figure 9.19—Transplant emergents early to avoid root malformation. This palm seedling was permitted to get too large in the 

tray, causing the root to grow at a 90° angle. This root malformation will cause poor growth and performance later in the plant’s 

life. Photo by Brian F. Daley.

Figure 9.20—Miniplugs are a viable option for growing seedlings that will later be transplanted to a larger container. Miniplugs work very well with species with very tiny seeds. Photo by 

Tara Luna.
• Keep seeds in place. This practice improves contact 

with the medium and minimizes the number of seeds 

washed out of the containers by irrigation or rainfall.

• Reflect heat when mulches are light colored, so seeds 

do not get too hot on bright, sunny days.

• Reduce the development of moss, algae, and liverworts (figure 9.22).

The recommended depth of the seed covering varies by 

species; a general rule is to cover the seed twice as deep as 

the seed is wide. If mulch is too shallow, seeds may float 

away in the irrigation water. If the mulch is too deep, small 

plants may not be able to emerge above it (figure 9.23).

Seeds requiring light need to be left uncovered. Very 

small seeds need to be left uncovered or barely covered with 

a fine-textured material such as fine-grade perlite or milled 

Sphagnum peat moss. Uncovered and barely covered seeds 

must be misted frequently to prevent them from drying 

out. After light-requiring and light-sensitive species have 

emerged and are well established, mulch can be applied to 

prevent moss and liverwort growth and to help keep the 

medium moist. 

Seed Germination and Sowing Options 179

Figure 9.22—Mulches help to prevent the development of 

mosses and liverworts, which can compete with the seedling. 

Photo by Thomas D. Landis.

Figure 9.23—A general rule of thumb for covering seeds with 

mulch is to cover the seed twice as deep as the seed is wide. Species requiring light for germination should never be covered 

with mulch, although mulch can be added after germination to 

reduce the growth of moss, liverworts, and weeds. Illustration 

by Jim Marin.

Figure 9.21—Seed mulches are important to hold the seeds in place and to 

moderate the surface temperature of the 

medium during germination. Common 

mulches include poultry grit (A, B) perlite (C), and vermiculite (D). Photos A 

and B by Craig R. Elevitch, and photos C 

and D by Thomas D. Landis.

B

A C

D
180 Tropical Nursery Manual

Germinating Fern Spores

Ferns (figure 9.24) and fern allies (moonworts, mosses, and horsetails) differ 

from seed plants in that they produce spores instead of seeds. An understanding of 

the life cycle of ferns is essential for successful fern propagation in nurseries.

Ferns have two life stages—the gametophyte and the sporophyte—the latter being 

the spore-producing fern plant with which we are all familiar. The sporangia (sporebearing structures) are variously placed on the lower surface of the leaves and occur in 

clusters known as sori. In many species, the sori are covered by specialized outgrowths 

of the leaf, known as the indusium, which lifts and shrivels when spores are ripe. A 

specialized layer of cells on the stalks of the spores, known as the annuli, contract and 

expand and the mature spores are disseminated with a catapult-like discharge.

After the spores disseminate, they germinate upon contact with a suitably moist 

substrate. Spore germination results in the gametophyte, which begins development 

as a small, pale-green, algae-like chain of cells known as the germ filament. Development continues into a flat, heart-shaped structure called the prothallus. Slender holdfasts, known as rhizoids, develop on the lower surface 

of the prothallus. The reproductive structures, the antheridia (male) and the archegonia (female), develop on the lower surface of the 

prothallus. Antheridia usually appear before the archegonia, mostly near the rhizoids. Archegonia appear near the notch of the prothallus.

Water must be present for the sperm to swim from the antheridia to the eggs in the archegonia. After fertilization, the young sporophyte 

receives its nutrients from the gametophyte via a foot-like structure. Further development is rapid and, after the sporophyte achieves a level 

of photosynthesis sufficient to maintain itself, the gametophyte disintegrates. The sporophyte completes the life cycle when it grows into a 

mature fern plant and produces spores.

Fern Propagation

When propagating ferns, maintaining a high level of sanitation during all phases of fern development is of the utmost importance. To 

collect fern spores, find fern fronds with ripe (dark brown, black, or gold) sori. Cut a piece off the frond. Lay each frond piece in a paper envelope, paper bag, or between two sheets of paper. Wait 24 to 48 hours for the spores to drop onto the paper, then collect (Lilleeng-Rosenberger 

2005). Removal of nonspore material reduces the chances of contamination by fungi, algae, or bacteria during spore germination. Spores 

can be sterilized by adding them to a 2- to 5-percent bleach solution with a small drop of wetting agent, which prevents the spores from 

sticking together. Soak for a few seconds to 1 minute. Collect spores on filter paper and rinse with distilled water thoroughly for 2 minutes 

(Hoshizaki and Moran 2001).

Germinate spores using sterilized propagation flats that are at least 2 in (5.2 cm) deep with drainage holes. You can use peat cubes, peat 

pellets, or a sterilized clay brick placed within an airtight, clear plastic shoebox container. Any sterilized commercial growing mix without 

added fertilizers can also be used. Moisten the medium thoroughly with distilled water before sowing.

Spores can be sown by spraying them on the medium surface with an atomizer, or by delivering small amounts of spores through a 

syringe with distilled water. Another option is dip a cotton swab into an envelope filled with spores, and then apply the spores to the growing 

media using the cotton swab.

After sowing, irrigate with distilled water and cover immediately with a clear plastic lid to seal in moisture and to prevent fungal contamination. Place flats under 150- to 500-foot candles of light. If you are using artificial light (cool, white fluorescent), leave lights on for 8 to 

24 hours per day. Optimum spore germination temperature is 68 to 86 °F (20 to 28 °C). Try to maintain a constant germination temperature 

to avoid excessive condensation in the sealed flats. Distilled water needs to be applied when the medium begins to dry slightly on the surface. 

Flats also need to be closely monitored for any fungal contamination. Spores germinate 10 to 20 days after sowing. If additional irrigation is 

needed, water only with distilled water delivered through a sterile mist bottle or by placing flats in trays filled with distilled water.

The presence of mold may require special treatments. Stop overhead watering. Make sure that water is not dripping excessively onto 

plants from condensation. Remove mold and at least 12 mm (0.5 in) of plant tissue and medium beyond the infected area. Apply a mild 

fungicide labeled for ferns if infection continues.

Shortly after spore germination, the thread-like germ filament can be seen with the aid of a microscope. In general, prothalli become 

visible 20 days after sowing. The prothalli continue to grow for up to 10 weeks before the reproductive structures, the antheridia and archegonia, become evident on the under surface of the prothallus. These structures can be seen with a microscope when sampling a few prothalli 

from a tray. After these structures appear, it is important to maintain a thin film of distilled water over the surface of the prothalli. It is very 

important to keep the germination surfaces evenly moist at all times.

The clear plastic lid is removed from the container when the antheridia have withered and disappeared, usually 4 weeks after their 

initial appearance. Flats are then transferred from under indoor lights to a shaded greenhouse. The young fern plants (sporophytes) with 

true leaves and a developing root system will appear sometime after fertilization; from a few weeks to a few months. They can then be 

transplanted into individual containers.

Figure 9.24—Native Hawaiian fern, a Sadleria

species. Photo by J.B. Friday.
Try Different Sowing Techniques 

and Keep Detailed Records

Native plant growers often work with seeds of species that have not yet been propagated in nurseries. Little 

literature or experience is available to answer questions 

about seed dormancy-breaking requirements, environmental requirements, germination percentages, and 

other factors. Understanding the biology and ecology of 

tropical plants will provide important clues on how to 

overcome seed dormancy (if any) and provide the correct environmental conditions needed for germination.

It is important to develop a good recordkeeping system to refine and improve results over time and prevent 

the loss of valuable information. Keep details on the general information of the species, seedlot, seed treatments, 

and resulting germination. To improve propagation 

results, use these details to develop and refine propagation protocols, as described in Chapter 4, Crop Planning: 

Propagation Protocols, Schedules, and Records.

Seed Germination and Sowing Options 181

Figure 9.25—Small trials of seed germination and sowing 

options will help you discover the most effective approach for different 

species. Photo by Brian F. Daley.

Trials To Develop a Successful Propagation Protocol 

for an Endangered Species

Catesbaea melanocarpa of Puerto Rico and the U.S. Virgin Islands is considered 

federally endangered throughout its range. The seeds (figure 9.26A) are small and challenging to work with.

A few individuals of Catesbaea melanocarpa were discovered in a pasture after 

a wildfire burned grass and underbrush that was obscuring them. With so few plants 

remaining and in danger of future fires, propagation by seed became a priority. After 

months of monitoring, a dozen or so ripe fruit were collected, each containing an average of eight small, flat seeds. The seeds were too small to be sown in traditional seed 

trays and monitored effectively. Due to their small size and scarcity, researchers at the 

University of the Virgin Islands decided to treat the seeds similar to the way they handle 

orchid seeds. Sterile dishes of sucrose rich agar (gel) were prepared and 6-8 seeds were 

placed in each of 10 dishes (figure 9.26B). This intensive germination method allowed 

for daily observations on every seed collected. Less than one-half of the seeds germinated, and later dissection revealed the nongerminated seeds were smaller than the germinated seed and did not have embryos. When the germinants produced adult leaves, 

the entire dish was transplanted into plant pots with a mix of ProMix (peat, perlite, and 

vermiculite blend) and coarse sand. More than 25 percent of the plants desiccated and 

died shortly after transplanting. The rest of the plants established, but grew slowly. Even 

with the addition of sand, the media seemed too coarse for the tiny plants.

Later in the season, more fruit were collected. The researchers now knew that 

one-half of the seeds were likely viable, but that the smaller, malformed seeds could 

be discarded because they do not have embryos. Researchers added vermiculite to the 

growing medium mix and sifted it through a screen. Seeds were sown directly into the 

new, fine medium and germinated in trays in the greenhouse with 80 percent success 

(figure 9.26C). The second set of seedlings grew more vigorously and did not suffer the 

transplant shock that killed plants in the first trial.

The first trial led to discoveries about the seeds’ viability and identified problems 

with the growing medium and transplant shock. The second trial used this information and resulted in high germination rates and healthy seedlings of a federally 

endangered plant in the greenhouse.

Figure 9.26—Seeds of Catesbaea melanocarpa shown here with a fruit, a fruit in cross 

section, and with cleansed seeds (A) were evaluated in sterile dishes (B) to develop a successful 

germination protocol for this endangered species 

(C). Photos by Brian F. Daley.

B

C

A
Because growers have a number of options for sowing seeds, it is a good idea to do small trials of several of 

the methods described in this chapter (figure 9.25). See 

Chapter 20, Discovering Ways to Improve Nursery Practices and Plant Quality, for proper ways of conducting 

trials. Although several methods may “work”—that is, 

result in a viable plant produced—the question during 

the trials should be: Which method is optimal? Trials will 

help you decide how to answer this question yourself.

Larger Role of Nurseries in 

Conservation of Species

Each seed is a link between the evolutionary processes 

of the past and the potential for future adaptation (Flores 

2002). While you clean, treat, and germinate seeds to grow 

your crops, be mindful how your actions affect that species genetic diversity and ability to adapt to the future. Do 

your best to maintain as much genetic diversity as possible 

within the species you propagate.

Nurseries are often involved in genetic selection beyond 

seed collection practices, intentionally or not. For example, 

production schedules may cause growers to favor faster germinating over slower germinating individuals of the same 

species, although the quality of the resulting plant would 

be similar. For some species, the earliest sprouters may be 

the healthiest. But in other cases, sprouting later may be an 

adaptive trait; one that could be selected out accidentally 

by nursery practices. Following up with plant performance 

in the field can reveal if, in fact, the slower germinating 

individuals grow well. If so, no reason exists to select these 

individuals out with nursery practices, and every reason to 

keep their traits in the gene pool. In this example, simply 

planting the slower sprouting individuals, as well as the fast 

ones, could protect diversity. This example is just one that 

shows how different steps in collecting, storing, germinating, and sowing seeds can affect subsequent plant genetics. 

The desire for uniform crop size and standardized schedule 

must be balanced with the need to protect and perpetuate 

species and genetic diversity.

Nurseries may also have an increasing role in the conservation of tropical species by helping protect and restore 

genetic diversity of recalcitrant species (species whose 

seeds do not store well). You are probably familiar with 

the concept of “seed banks”—seed storage facilities used as 

reserves to protect and restore species in case their habitats 

are threatened. Seed banks are also used for some traditional food crops that have become rare with conventional 

agriculture. Seed banks work well for orthodox species, 

whose seeds can store for many years and remain viable. 

In tropical ecosystems, however, recalcitrant species are as 

numerous and as important as orthodox species. Because of 

this distinction, many tropical species cannot be “banked” 

in conventional seed banks. Instead, nurseries and seedling 

propagation efforts will play key roles in any efforts to conserve and restore tropical recalcitrant species (Kettle and 

others 2011).

References

Baskin, C.C.; Baskin, J.M. 1998. Seeds: ecology, biogeography and 

evolution in dormancy and germination. San Diego, CA: Academic Press. 666 p.

Baskin, C.C.; Baskin, J.M. 2004. Determining dormancy-breaking 

and germination requirements from the fewest numbers of seeds. 

In: Guerrant, E.O., Jr.; Havens, K.; Maunder, M., eds. Ex situ plant 

conservation: supporting species survival in the wild. Washington, 

DC: Island Press: 162–179.

Drake, D.R. 1993. Germination requirements of Metrosideros polymorpha, the dominant tree of Hawaiian lava flow and rainforests. 

Biotropica. 25: 461–467.

Dumroese, R.K.; Landis, T.D.; Wenny, D.L. 1998. Raising forest 

tree seedlings at home: simple methods for growing conifers of 

the Pacific Northwest from seeds. Contribution 860. Moscow, ID: 

Idaho Forest, Wildlife and Range Experiment Station. 56 p.

Dumroese, R.K.; Wenny, D.L.; Quick, K.E. 1990. Reducing pesticide use without reducing yield. Tree Planters’ Notes. 41(4): 

28–32.

Flores, E.M. 2002. Seed biology. In: Vozzo, J.A., ed. 2002. The 

tropical tree seed manual. Agriculture Handbook 721. Washington, DC: U.S. Department of Agriculture, Forest Service. 13–118. 

Chapter 1.

Gao, Y.P.; Zheng, G.H.; Gusta, L.V. 1998. Potassium hydroxide 

improves seed germination and emergence in five native plant 

species. HortScience. 33: 274-276.

Hartman, H.T.; Kester, D.E.; Davies, F.T.; Geneve, R.L. 1997. 

Plant propagation: principles and practices. 5th ed. Upper Saddle 

River, NJ: Prentice Hall Press. 770 p.

Hong, T.D.; Ellis, R.H. 2002. Storage. In: Vozzo, J.A., ed. 2002. 

The tropical tree seed manual. Agriculture Handbook 721. 

Washington, DC: U.S. Department of Agriculture, Forest Service. 

13–118. Chapter 3.

Hoshizaki, B.J.; Moran, R.C. 2001. The fern growers manual. 

Portland, OR: Timber Press. 604 p.

182 Tropical Nursery Manual
James, R.L.; Genz, D. 1981. Ponderosa pine seed treatments: 

effects on seed germination and disease incidence. Report 81-16. 

Missoula, MT: U.S. Department of Agriculture, Forest Service, 

Northern Region, Forest Pest Management. 13 p.

Keeley, J.E.; Fotheringham, C.J. 1998. Smoke induced seed 

germination in California chaparral. Ecology. 79: 2320-2336.

Kettle, C.J.; Burslem, D.F.R.P.; Ghazoul, J. 2011. An unorthodox 

approach to forest restoration. Science. 333: 35. 

Kurbitzky, K.; Ziburski, A. 1994. Seed dispersal in floodplain forests of Amazonia. Biotropica. 26: 30–43.

Landis, T.D.; Tinus, R.W.; Barnett, J.P. 1999. The container tree 

nursery manual: volume 6, seedling propagation. Agriculture 

Handbook 674. Washington, DC: U.S. Department of Agriculture, Forest Service. 166 p.

Lilleeng-Rosenberger, K. 2005. Growing Hawaii’s Native Plants. 

Honolulu, HI; Mutual Publishing. 420 p.

Mousa, H.; Margolis, H.A.; Dubay, P.A.; Odongo, J. 1998. Factors 

affecting germination of doum palm (Hyphaene thebaica Mart.) 

from the semi-arid zone of Niger, West Africa. Forest Ecology 

and Management 104 (1-3): 27-41.

Narimanov, A.A. 2000. Presowing treatment of seeds with hydrogen peroxide promotes germination and development in plants. 

Biologia. 55: 425–428.

Vozzo, J.A., ed. 2002. The tropical tree seed manual. U.S. Department of Agriculture, Forest Service. Washington, DC: U.S. Government Printing Office. 899 p.

Additional Reading

Dreesen, D. 2004. Tumbling for seed cleaning and conditioning. 

Native Plants Journal. 5: 52–54.

Koebernik, J. 1971. Germination of palm seed. International Palm 

Society. 15: 134–137.

Landis, T.D. 2000. Where’s there’s smoke...there’s germination? 

Native Plants Journal. 1: 25–29.

Schmidt, L. 2007. Tropical forest seed. Berlin, Germany: SpringerVerlag. 409 p.

Seed Germination and Sowing Options 183
184
185

Many desirable and ecologically important tropical plant species can be difficult or 

very time consuming to propagate by seeds. Thus, nursery growers may want to investigate how to propagate these species by vegetative propagation, which is accomplished 

by combining classic horticultural propagation techniques with an understanding 

of the ecological and reproductive characteristics of the species. Plants that inhabit 

tropical ecosystems often reproduce vegetatively (that is, asexually without seeds or 

spores). Vegetative propagation is commonly used with species that have short seed 

life, low seed viability, or complex seed dormancy. All new daughter plants that arise 

from vegetative propagation are genetically identical to the mother (donor) plant, and 

these resulting individuals are known as “clones” (figure 10.1).

Vegetative Propagation

Tara Luna and Diane L. Haase 10

Facing Page: Vegetative propagation is the production of daughter plants from the stems, leaves, roots, or other portions of a 

single mother (donor) plant. Photo by Ronald Overton.
Comparing Vegetative to Seed 

Propagation

The following situations favor vegetative propagation 

over seed propagation:

• Seed propagation is difficult or very time-consuming.

• Viable seeds are produced infrequently or in 

small quantities.

• Larger nursery stock is needed in a shorter 

period of time.

• An individual, unique plant needs to be propagated.

• A need exists to shorten time to flower for 

seed production.

• A uniform stocktype is needed.

• Specific genotypes are desired.

• Disease-free nursery stock is required.

Disadvantages of using vegetative propagation include—

• Greater production costs than seed propagation, 

usually because of increased labor.

• Reduced genetic diversity.

• Specialized propagation structures may be 

required, based on the time of year and the species 

to be propagated.

Depending on the plant species, vegetative propagation can be done with pieces of stems, leaves, roots, bulbs, 

corms, tubers, and rhizomes. The species, the type of vegetative material used, the time of year that material is collected, how it is handled and manipulated to induce rooting, and proper application of the correct environmental 

conditions all affect vegetative propagation.

Because vegetative propagation is more costly than 

growing seedlings, the production system must be efficient. 

A general rule of thumb is that at least 50 percent rooting 

must be obtained to produce cuttings economically. If rare 

species or individual plants are being propagated, however, 

costs may be less important. 

Consider these methods to reduce production costs:

• Develop a smooth production line, from the 

collection of material to the final product.

• Train nursery staff how to properly collect, process, 

plant, and grow material.

• Build a dibble for making holes in the rooting 

medium.

• Keep good records to improve your results and to 

document production costs.

Cuttings

A cutting is the portion of a plant that is collected, treated, 

and planted to develop into a new intact plant complete with 

stems, leaves, and roots. Cuttings can be collected from mother 

plants in the wild, or special donor plants can be cultured in 

the nursery. Selection of mother plants must be done carefully; 

it is as important as the origin of seeds to ensure that nursery 

stock is well adapted to the outplanting environment. Collection of cuttings should follow the same ethical guidelines as 

collection of seeds to establish proper genetic diversity and 

sustainability of wild populations (see Chapter 8, Collecting, 

Processing, and Storing Seeds). In addition, the ability of cuttings to root is often clone-specific, so it is important to record 

the origin of cuttings and subsequent rooting success.

Striking is the process of placing the cutting into soil or 

a rooting substrate. Often, propagators will say that cuttings have been “struck” to indicate that the cuttings have 

been placed in the rooting substrate.

Shoot or Stem Cuttings

Shoot cuttings, also referred to as stem cuttings, are the 

most common type of cuttings and can be broadly placed 

186 Tropical Nursery Manual

Figure 10.1—Propagation of plumeria by cuttings produces 

genetically identical plants. Photo by Tara Luna.
into three categories depending on the stage of growth they 

are in when collected. Hardwood cuttings are made from 

the previous year’s mature wood of trees and shrubs and 

are usually collected during the dry season in the tropics, 

when the leaves of deciduous species have dropped (figure 

10.2). Semihardwood (greenwood) cuttings are collected 

towards the end of the active growth season when stem tissues have hardened and terminal buds have formed or just 

after a flush of growth has taken place and the wood is partially matured. Softwood cuttings are collected when stems 

and leaves are actively growing.

Hardwood Cuttings of Deciduous Species

Tropical deciduous hardwood stem cuttings are the easiest, least expensive type of cuttings because they are easy 

to prepare, are not as perishable as softwood or semihardwood cuttings, can be stored in coolers or shipped if necessary, and require little or no special equipment during rooting. They are sometimes struck directly on the outplanting 

site or brought back to the nursery to grow as bareroot or 

container stock.

If deciduous hardwood cuttings are struck directly on 

the outplanting site, they can be live stakes (12- to 16-in 

[30- to 40-cm] long), poles (12- to 16-ft [3.6- to 4.9-m] 

long), or branched cuttings (2- to 6-ft [0.6- to 1.8-m] long). 

These cuttings are collected during the dry season until 

the early onset of the rainy season and outplanted when 

the cutting is leafless and the soil at the outplanting site is 

wet. Live stakes and branched cuttings are usually driven 

into the ground with a mallet and need to be planted deep 

enough to reach moisture in the soil profile with only three 

to four nodes (buds) above ground. Poles are much longer 

and are also driven deep enough so they can be in contact 

with the water table, but the aboveground stem is much 

longer. Hardwood pole cuttings of Erythrina, Gliricidia,

and other species are commonly used this way for windbreaks, living fences, and restoration projects.

If hardwood cuttings are struck in the nursery, they can 

be straight, heel, or mallet cuttings (figure 10.3). Straight 

cuttings are made from straight hardwood stems and are 

the most common type for easy-to-root species. Heel cuttings are made from 2-year-old side shoots. To make a heel 

cutting, pull the side shoot away from the tip so that a section of older wood remains at the base of the cutting. Mallet 

cuttings include a cross-section of older stem at the base of 

the side shoot.

All hardwood stem cuttings have an inherent polarity 

and will produce shoots on the distal end (nearest the bud) 

and roots on the proximal end (nearest the main stem or 

root system). If planted upside down, the cutting will not 

root. When using straight or live stake deciduous cuttings, 

the tops and bottoms of the stems need to be distinguished. 

The absence of leaves can make it difficult to discern the 

top from the bottom, so it is useful to cut the bottoms diagonally and the tops straight across. The diagonal cut maximizes water uptake area at the base, and the straight cut 

minimizes the water loss area at the top.

Vegetative Propagation 187

Figure 10.2—Hardwood cuttings are collected during the dry season or when deciduous tropical species shed their leaves. Photo 

by Thomas D. Landis.
Hardwood Cuttings of Evergreen Species

Hardwood cuttings of broadleaf tropical evergreen and 

coniferous species are usually taken during the dry season 

or just after a new flush of growth during the growing season. Unlike deciduous hardwood cuttings, evergreen cuttings must be struck into a special rooting environment (see 

Chapter 5, Propagation Environments) as soon as possible because they cannot be stored for any length of time. 

Evergreens are best rooted in special rooting environments 

after being wounded or treated with rooting hormone 

(described in the following sections). Cuttings are usually 

4 to 8 in (10 to 20 cm) long, with all leaves removed from 

the lower half. Green tips and side shoots also need to be 

removed. The large leaves of tropical broadleaf evergreen 

plants are usually cut in half to reduce water loss during 

rooting (figure 10.4). Straight, mallet, and heel cuttings are 

also used with evergreen species (figure 10.3).

Semihardwood Cuttings

Semihardwood (greenwood) stem cuttings are made from 

newer shoots of leafy broad-leaved evergreen plants and leafy 

deciduous species. Cuttings are taken just before the onset 

of the dry season, towards the end of the active growth season when stem tissues have hardened, or just after a flush of 

growth when the wood is partially matured. In many cases, the 

terminal bud has formed. Semihardwood cuttings are propagated in the same manner as evergreen hardwood cuttings.

Softwood Cuttings

Prepared from the new growth of deciduous or evergreen species, softwood cuttings generally root easier than 

other types of cuttings but require a special rooting environment and more attention to prevent desiccation. The 

best cutting material has some degree of flexibility but is 

mature enough to break when bent sharply (figure 10.5). 

Extremely fast-growing tender shoots are not desirable.

Herbaceous stem cuttings are softwood cuttings made 

from nonwoody plants. They are handled in the same way as 

softwood cuttings (figure 10.6). Many succulent tropical plant 

cuttings, including some cacti, are easily propagated in this 

manner; cuttings need to be allowed to develop callus for a 

week before inserting the cutting into rooting media. Succulent cuttings root readily without misting or high humidity.

Root Cuttings

Although not used as much as other types of cuttings, 

root cuttings can be made by dividing roots into individual 

segments containing dormant shoot buds capable of developing into new plants. Root cuttings are typically used on 

species that fail to root well from stem cuttings. Breadfruit 

(Artocarpus species) and noni (Morinda citrifolia) are commonly propagated from root cuttings (figure 10.7). Root 

sections are collected any time of the year in the tropics.

188 Tropical Nursery Manual

Figure 10.4—The leaves of some broadleaf evergreen cuttings 

are usually cut in half to reduce the amount of water loss during 

rooting. Photo by Thomas D. Landis.

Figure 10.5—Rooted softwood stem cuttings of hibiscus. Softwood stem cutting material has some degree of flexibility but is 

mature enough to break when bent sharply. Photo by Tara Luna.

Figure 10.3—Left to right: straight, heel and mallet cuttings. 

Straight cuttings are used on easy-to-root species, while mallet 

and heel cuttings are used on species that are more difficult to 

root. Photo by Tara Luna.
Root cuttings are planted horizontally in planting beds 

or containers with the dormant leaf buds on the upper side. 

Some root cuttings are also planted vertically, but it is important to maintain the correct polarity. Root cuttings generally 

do not require a special rooting environment unless shoots 

are cut from the root piece and treated as a stem cutting.

Selecting Cuttings From 

Mother Plants

A variety of factors such as seasonal timing, juvenility, 

plagiotropism, species, and cutting size and quality can 

greatly influence the rooting success of cuttings. Collectors 

need to be aware of these factors and, with experience, 

will be able to discern the right type of cutting material 

to collect.

Some species can be readily propagated from cuttings 

collected in any season of the year, while others have 

very specific seasonal trends when they will form roots. 

For any given species, small experiments are required to 

determine the optimum time to take cuttings, which is 

related to the physiological condition of the donor plant 

at collection time rather than any given calendar date. 

Recordkeeping is important to improve rooting results 

from year to year.

All plants progress from a juvenile phase (incapable of 

producing flowers) to a mature or adult flowering phase. Different parts of the plant, however, can be at different stages 

of maturity at the same time. Sometimes the juvenile phase 

can be distinguished from the adult phase by differences in 

leaf shape or color or by the overall habit of the plant. Some 

broadleaf species, such as eucalyptus, have distinct juvenile 

leaves that differ from adult leaves. In conifers, juvenile wood 

is usually found on the lower portion of the tree crown and 

the adult, cone-bearing wood is located in the upper crown. 

In broadleaf plants, juvenile wood is found near the stem base 

or root crown and can be discerned as the long, nonflowering 

shoots (sucker shoots). Cuttings collected from this region 

of the plant root more easily than those from older, mature 

wood. In some cases, many difficult-to-root species will root 

only from stems collected from young seedlings. Hedging 

or coppi cing is the practice of regularly cutting back donor 

plants to maintain juvenile wood and is an efficient means of 

generating many long, straight cuttings from a limited number of plants. Donor plants in natural stands can be selected 

for hedging on an annual basis if cuttings will be collected 

from the area for several years. Otherwise, mother plants 

can be held in the nursery and used as a source of cuttings as 

described in the next section.

Plagiotropism is the habit of a cutting to continue to 

grow in the direction it was growing on the donor plant. 

Plagiotropism can be strong or weak depending on species and on the original position of the cutting on the 

donor plant. Plumeria is an example of a tropical broadleaf genus with strong plagiotripism. Often, plants produced from cuttings from lateral shoots will maintain a 

lateral habit, whereas plants produced from terminal 

shoots will grow vertically. This tendency can create 

problems with the growth habit of the nursery stock and 

is more of a concern with conifers than broadleaf species 

(Landis and others 1999) (figure 10.8).

Vegetative Propagation 189

Figure 10.7—Root cuttings, such as those sown here from 

breadfruit (Artocarpus species), can be used when stem cuttings 

do not root well. Photo by Thomas D. Landis.

Figure 10.6—Many herbaceous tropical species, such as 

āwikiwiki (Canavallia species) can easily be rooted from cuttings 

using the application of a rooting hormone and a mist environment or an enclosed poly propagator. Photo by Tara Luna.
Cutting size varies from species to species and by cutting type and seasonal timing. Easily rooted plants can be 

collected as long poles for rooting or made into small microcuttings. Microcuttings consist of one bud and a small section of internode stem and are typically less than 2 in (5 cm) 

long (figure 10.9). Hardwood cuttings vary in length from 4 

to 30 in (10 to 76 cm). At least two nodes are included in the 

cutting. The basal cut is made just below a node and the top 

cut is made above a node. If more than one cutting is being 

made from a stem, be certain that nursery workers maintain 

the correct polarity. The very tip portions of the shoot, which 

are usually low in carbohydrates, are usually discarded. Central and basal portions of the stem usually make the best 

cuttings, but exceptions exist. Good cutting wood has some 

stored carbohydrates that will supply the cutting with food 

reserves until roots form. Very thin or elongated shoots are 

not desirable. If cuttings are collected from natural stands, 

harvest from individuals that are growing in full sun to 

partial shade and avoid those in deep shade. Often, the ability of a cutting to produce new roots changes from the base 

of the cutting to the tip. Softwood stem cuttings are usually 

straight, 3 to 6 in (7.5 to 15 cm) long with two or more nodes. 

In general, softwood cuttings root better from terminal 

shoots. Semihardwood cuttings are usually 3 to 6 in (7.5 to 

15 cm) long with leaves retained in the upper end. Semihardwood cuttings usually root best from lateral shoots.

Many tropical plant species are dioecious, meaning 

that male and female flowers are borne on separate plants 

(table 10.1). For example, Hawai‘i has the highest degree 

of diocity in its native flora than anywhere in the world. 

In such cases, collectors may not realize they have collected cuttings of only one sex. Outplanting plants of only 

one sex onto the restoration site may compromise project 

objectives because seed production over the long term will 

be impossible. Therefore, be sure to collect both male and 

female cutting material (see Landis and others 2003).

Establishing Mother Plants 

at the Nursery

Some nursery managers find it advantageous to maintain donor stock plants at the nursery as a continual 

source of cutting material in a convenient location. Most 

broadleaf tree species will coppice (regenerate stems from 

cut stumps), which can provide new cuttings year after 

year. Maintaining stock plants at the nursery can be more 

efficient than collecting from wild populations, especially 

if the same ecotypes will be used for a long-term restoration project. The disadvantage to using mother plants 

grown at the nursery is that they require nursery space 

and must be intensively managed.

Mother plants are usually planted in field beds at 

the nursery or, in some cases, are kept in large containers. These areas are often referred to as “stooling beds” 

190 Tropical Nursery Manual

Figure 10.9—Microcuttings are small stem cuttings with one 

or two nodes. Photo by Tara Luna.

Table 10.1—Tropical dioecious species.

Scientific name Common name

Aleurites moluccana Kukui nut

Broussaisia arguta Kanawao

Cycas species Cycad

Morus species Mulberry

Pandanus tectorius Podocarp

Piper methysticum Kava

Pittosporum hosmeri Ho`awa

Zanthoxylum flavum and Z. thomasianum Yellow prickle

Figure 10.8—Plagiotropism is the effect of the position of the 

branch on the growth habit of the progeny, The terminal shoot on the 

juniper cutting on the right was collected from a lateral branch and 

still exhibits lateral growth tendency. Photo by Thomas D. Landis.
or “hedgerows.” Mother plants must be clearly labeled as to 

species and origin. If mother plants are in field beds, an accurate map should be kept. Mother plants should be hedged 

on an annual basis to maintain wood juvenility, discourage 

thick shoots or dominant leaders, and encourage production of numerous straight shoots to use as cutting material. 

When hedging, it is important to leave enough leaves to keep 

the root system alive. Moderate shade will encourage shoot 

elongation, thereby resulting in longer internodes and more 

easily rooted cuttings (Longman 1993). Mother plants also 

need periodic watering and nutrients and should be kept free 

of weeds by mulching or other means.

Collecting, Transporting, and 

Storing Cuttings

Some basic equipment and supplies are necessary to 

efficiently collect cuttings and ensure their health until 

they are struck (figure 10.10). The following items are 

recommended:

• High-quality, sharp pruning shears and pruning 

poles for collecting from trees.

• Spray bottles filled with disinfectant (1 part bleach 

[5.25 percent sodium hypochlorite] in 10 parts water) 

to disinfect pruning shears.

• Permanent labels and marking pens for noting 

origin of collection.

• Large, white plastic bags with ties for bulk collections.

• Spray bottles filled with water to keep cuttings 

moist in the plastic bags after collection.

• Portable, insulated coolers for transport back 

to the nursery.

• Newspaper, moss, or other materials to moisten 

and wrap around cuttings.

When collecting and handling cuttings, it is important to—

• Collect only from healthy donor plants.

• Keep cuttings cool to avoid wilting and desiccation.

• Handle cuttings carefully so that tissues are 

not bruised.

• Make sure that some buds or leaves are present on 

stem cuttings.

• Collect from nonflowering shoots. In general, 

cuttings root better before or after flowering.

• Place cuttings in the same direction when bundling 

to avoid mix-ups with polarity.

Cuttings should be collected on cloudy, cool days or 

during the early morning. All cuttings should be handled 

with care. Cuttings need to be kept cool and shaded during collection and transport back to the nursery to avoid 

water loss and physical damage. Wrap cuttings loosely 

in moistened newspaper (or other moistened material to 

protect from desiccation), place them into white plastic 

bags, mist them, and label with origin information and the 

date. When collecting from mother plants, make a proper 

cut that facilitates healing of the mother plant. Take the 

cutting just above a node, ensuring that you do not leave 

a stub. Then trim the base of the cutting to just below the 

node where rooting is more likely to occur. Between collection sites, disinfect the pruning shears with the bleach 

solution to avoid spreading disease.

Deciduous hardwood cuttings can be wrapped in moist 

peat moss or burlap and stored in a shaded, dry environment 

with periodic moisture to prevent dessication. Cuttings can 

be stored for several days but generally no longer than 4 to 

8 weeks. Inspect stored cuttings frequently to make certain 

that tissues are slightly moist and free from fungal diseases. 

Hardwood and softwood evergreen cuttings, deciduous softwood cuttings, and semihardwood cuttings should be struck 

in propagation beds the same day of collection and should 

never be stored for longer than 1 day.

Types of Rooting

The development of new roots on a shoot is known as 

“adventitious root formation” (figure 10.11A). Two types of 

roots occur depending on whether buds capable of producing new roots are present. Many tropical species have buds 

present and the resulting roots are termed “preformed” or 

“latent.” In the nursery, cuttings of these species are usually 

struck directly into containers because they do not require a 

special rooting environment. This method is the easiest and 

most economical way to produce these species because no 

additional transplanting is needed.

Vegetative Propagation 191

Figure 10.10—Equipment used to collect cuttings includes 

sharp tools, a cleaning agent for tools, and a cooler to keep cuttings 

from drying out during transport. Photo by Tara Luna.
If no buds are present, then the roots are termed “woundinduced” and new roots will form only in response to the 

wound caused by preparing the cutting. Species requiring 

wounding can vary considerably in their ability to form new 

roots. After a cutting is wounded, callus tissue forms at the 

base, primarily from the vascular tissue (figure 10.11B). In 

easy-to-root species, callus formation and root formation are 

independent processes that occur at the same time because 

of similar environmental triggers (figures 10.11B, 10.11C). 

In difficult-to-root species, adventitious roots arise from the 

callus mass. In some cases, excessive callus can hinder rooting and is a signal to use a lower concentration of rooting 

hormone. Often, excess callus needs to be scraped away and 

the cutting replaced in the rooting environment.

In general, all species with wound-induced roots must 

first be rooted in a special propagation environment with 

tightly controlled air and medium temperatures, high relative humidity, reduced light levels, and “moist, but not wet” 

medium (figure 10.12). See Chapter 5, Propagation Environments, for more details on propagation environments. Easyto-root species are often struck directly into containers filled 

with regular growing medium and, after they are rooted in 

the special propagation environment, they are moved into 

the regular nursery. Hard-to-root species are often struck 

into trays or beds containing a special rooting medium and, 

after roots form, they are transplanted into containers to 

continue their growth. 

Cutting Preparation

While preparing cuttings, it is important to keep the work 

area clean. Use sharp, well-maintained shears and knives to 

make clean cuts. Disinfect these tools often to reduce the 

possible spread of disease. Preparing cuttings standardizes 

their size and shape, promotes side shoots, and eliminates 

shoot tips that often die back. It is important to maintain 

polarity during this process, especially for deciduous hardwood cuttings. Cuttings that will require hormone treatment 

to encourage rooting, such as those of hardwood narrowleaf 

evergreens or any softwood or semihardwood cuttings, 

should have one-third to one-half of the leaves and buds 

removed to reduce the amount of water loss from the cutting. Any flower buds also need to be removed. It is important, however, to retain some buds or leaves on the cutting so 

that the cutting can manufacture food during rooting.

Wounding Cuttings

Wounding, used on species that are difficult to root, 

increases rooting percentages and improves the quantity 

and quality of roots produced. Wounding exposes more 

cells to rooting hormone, encourages callus formation, 

and, in some cases, removes thick woody tissue that can 

be a barrier to root formation. Cuttings are commonly 

wounded by hand-stripping small lower stems and leaves 

to create wounded areas along the basal portion of the cutting, scraping the base of the stem with a small, sharp knife 

192 Tropical Nursery Manual

Figure 10.12—Semihardwood cuttings in a shadehouse at the 

University of the Virgin Islands, agroforestry facility are struck in 

trays of 100 percent perlite and misted every hour to ensure that 

no standing water exists, but the cuttings are always moist. The 

shadecloth around the PVC frame provides a shady, low-wind, 

high-moisture atmosphere for the cuttings while they produce 

roots. Rooted cutting are later transferred to small pots in the shadehouse for 2 to 3 weeks and later transferred to full sun. Photo by 

Brian F. Daley.

Figure 10.11—Adventitious roots of a cutting (A), callus and roots forming at the base of a cutting (B), adventitious root formation during a 6-week period (C). Photos by Tara Luna.

A B C
or potato peeler (figure 10.13), or slicing one or two long, 

shallow slivers (0.75 in to 1.25 in [2 to 3.2 cm] long) of tissue from the base of the stem, making sure to penetrate the 

cambium layer of the stem. Slicing requires precision and 

experience so that cuttings are not excessively damaged.

Rooting Hormones

Auxins are natural plant hormones that encourage 

root formation on cuttings and are available from natural 

and synthetic sources. In practice, auxins are commonly 

referred to as rooting hormones. Most cuttings are treated 

with synthetic hormones that are available in powder and 

liquid form, and some preparations may contain chemical fungicides (figure 10.14). Synthetic hormones can be 

purchased ready to use or can be mixed by growers to 

specific concentrations using ingredients purchased from 

horticultural suppliers. Indole-3-butyric acid (IBA) and 

naphthaleneacidic acid (NAA) are the most widely used 

synthetic auxins. Often, mixtures of IBA and NAA are 

more effective than either component alone. The effect of 

rooting hormones varies widely between species and, in 

some cases, between genotypes. In general, rooting hormone powders are expressed as a percentage, while liquid 

solutions are expressed as parts per million (ppm).

It is generally easiest to purchase ready-to-use formulations. It is important to remember that all rooting hormones 

have a limited shelf life of 18 to 24 months. Therefore, when 

purchasing or mixing hormones—

• Record the date of purchase on the container.

• Order only what you plan to use within 18 to 24 

months. Order smaller quantities more often to ensure 

that the rooting hormone remains effective.

• Keep containers sealed and refrigerated when not in use 

to preserve the activity of the rooting hormone.

Many growers prefer powders because several commercial products of varying strengths are available, they are easy 

to use, and large quantities of cuttings can be treated quickly. 

Powder must be applied uniformly to all cuttings; variable 

amounts of rooting powder adhere to the base of a cutting, 

which can affect rooting results (figure 10.15). The following 

precautions and special techniques are necessary when using 

powders:

• Wear gloves during application.

• Transfer enough hormone to a smaller container from 

the main stock container for use; never transfer unused 

hormone back to main stock container.

• Make sure the base of the cutting is moist so that the 

powder adheres; pressing cuttings lightly onto a moist 

sponge is a useful technique.

Chapter Title 193

Figure 10.14—Advantages of rooting hormones are (1) increased rooting percentages if applied correctly at an effective 

concentration, (2) more rapid root initiation, (3) an increase 

in the total number and quality of roots, and (4) more uniform 

rooting. Photo by Tara Luna.

Figure 10.13—Wounding the lower end of a stem cutting increases rooting with difficult-to-root species. Photo by Tara Luna.

Figure 10.15—Many growers prefer powders to liquid hormones 

(A). Apply powder hormones evenly and consistently (B). Photo A 

by Thomas D. Landis, and photo B by George Hernández.

A B
• Apply the hormone uniformly. Make sure cuttings are 

dipped into the powder to a depth of 0.2 to 0.4 in (5 

to 10 mm). Make certain that cut surfaces and other 

wounds are also covered with rooting hormone.

• Remove excess powder by lightly tapping the cuttings 

on the side of the dish.

Liquid products are formulated with alcohol and 

often must be diluted with great care to create the desired 

strength. Some of the advantages of using solutions are 

the availability of a wide range of commercial preparations, specific concentrations can be formulated at the 

nursery, and they can be stored for longer periods under 

the right conditions. Some growers believe that liquid 

solutions are more accurate than powders regarding the 

amount of rooting hormone entering the stem tissue. The 

most common procedure for treating cuttings with liquid 

products is the quick-dip method in which the base of the 

cutting is dipped into the solution for 3 to 10 seconds. 

Whole bundles of cuttings can be treated at once (figure 

10.16). An alternate method is to soak cuttings for a longer time in a more dilute hormone solution. When using 

liquid rooting hormones, it is important to—

• Wear gloves during mixing, preparation, and 

application.

• Make certain that the solution was precisely diluted to 

the correct concentration.

• Place the solution in a clean jar.

• Ensure that the treatment time is constant for a 

uniform application rate and to avoid damaging the 

plant tissue.

• Make certain that the basal ends are even to 

obtain uniform depth of dipping in the solution if 

bundles of cuttings are dipped.

• Allow the alcohol to evaporate from the stem of 

the cutting before striking cuttings into the propagation bed, a process that usually takes only a couple 

of minutes.

• Properly discard any remaining solution, because 

it is contaminated with plant material.

The optimum auxin rate for cuttings varies by plant 

species. A good starting rate is a 0.25-0.5 percent (2,500-

5,000 ppm). With trial and error, this rate can be increased 

or decreased until optimum rooting occurs for a particular 

species. Longman (1993) listed rates of IBA for the following tropical species:

• 0.2 percent for Triplochiton scleroxylon, Tripvochysia 

hondurensis, and several other tropical tree species.

• A range of 0.05 to 0.4 percent for Albizia guachapele.

• 0.4 percent for Cordia alliodora.

• 1.0 percent for Khaya ivorensis.

Striking, Monitoring, and 

Growing Cuttings

Direct striking into containers is more efficient and 

therefore more economical than striking into a special 

rooting environment because the cuttings are handled 

only once and expensive transplanting is avoided. Easyto-root hardwood cuttings, such as many Erythrina species, Gliricidia sepium (known locally as “quick stick” in 

much of the world), and mangrove propagules should 

always be direct struck (figure 10.17). Often, a dibble of 

the same diameter as the stem of the cutting is a useful 

tool for making openings in the medium into which the 

cutting can be struck. For striking several small cuttings, 

a template for the holes can be created by driving nails 

through a piece of plywood at the correct spacing and 

194 Tropical Nursery Manual

Figure 10.16—Using the liquid hormone “quick dip” method is 

preferred by many growers because bundles of cuttings can be treated 

at the same time with consistent uniformity of application. Photo by 

Tara Luna.
depth. If using powdered rooting hormones, this practice 

of creating holes ahead of time will help keep the hormone from being brushed off. The following practices 

need to be encouraged when striking cuttings:

• Wear gloves if the cuttings were treated with rooting 

hormones.

• Maintain polarity (keep the 

correct end of the cutting up).

• When using stem cuttings, make certain that at least two 

nodes are below the surface of the rooting medium.

• If cuttings were wounded, make certain that wounded 

tissue is adequately covered with rooting hormone and 

is below the surface of the rooting medium.

• Strike cuttings firmly in the rooting medium. Make 

certain to avoid air pockets around the base of the stem.

• Try to strike cuttings within 1 to 2 days so that all the 

plants will have the same level of root development and 

thus can be hardened off properly before harvesting.

• Place labels before the first cutting and after the final 

cutting of a particular clone or batch of cuttings.

After cuttings are struck, maintain a clean rooting 

environment (figure 10.18); routinely inspect cuttings for 

proper temperature, humidity, and moisture levels and 

adjust as necessary. Check to ensure that all equipment 

(including bottom heat) is working properly.

Environmental Conditions for Direct 

Struck Cuttings

In general, easy-to-root hardwood cuttings directly 

struck into containers can be treated similar to seedlings.

Vegetative Propagation 195

Figure 10.18—The propagation environment must be carefully 

maintained to ensure that the mist system, the timers that control 

the frequency of mist, and other equipment are working properly. 

Keeping the rooting environment as clean as possible during rooting 

is crucial for producing healthy plants. Photo by Tara Luna.

Environmental Conditions in Special 

Rooting Environments

Achieving successful rooting requires attention to sanitation, relative humidity, temperature, light, rooting medium, 

and sometimes mycorrhizae and dilute fertilization. See 

Chapter 5, Propagation Environments, for information 

about equipment necessary to regulate humidity, temperature, and light.

Sanitation

Always keep the propagation environment as clean as 

possible. Strike cuttings into a sterilized rooting medium. 

Routinely inspect for and remove dead leaves or cuttings 

that could be a source of disease infection.

Figure 10.17—Easy-to-root hardwood cuttings (A) or mangrove propagules (B) can be directly stuck into containers for rooting 

and is the most economical way of producing cuttings. Photo A by Tara Luna, and photo B by Thomas D. Landis.

A B
Humidity

Until the root system forms, high relative humidity 

must be maintained in the propagation environment to 

slow the rate of water loss from the cutting. Placing cuttings under an automatic misting or fogging system is 

often effective. Another method is to place the cuttings 

within a frame covered in clear or white polyethylene 

sheeting with a reserve of water below a moist rooting 

medium (Longman 1993). Cuttings in this “polypropagator” need to also be irrigated with a fine spray of water 

in late afternoon and early morning, especially when the 

weather is hot and dry. If possible, water temperature 

should not be significantly cooler or warmer than the 

rooting medium to avoid damaging the young roots from 

sudden changes in temperature. Achieving optimum 

humidity and medium moisture can be one of the most 

challenging aspects of successful propagation with cuttings. Too much moisture can encourage fungal pathogens and rotting, whereas too little moisture can result in 

lethal desiccation. Daily monitoring is important.

Light and Temperature

Providing light for photosynthesis is necessary so that 

cuttings can continue to manufacture food during rooting, but too much sunlight can cause high temperatures. 

Shadecloths of 30 to 50 percent shade cover are most effective to reduce air temperature while providing sufficient 

light. Shadecloth can also serve to reduce the effect of rain 

when propagating outdoors. The optimum air temperature 

for rooting cuttings is 68 to 80 °F (20 to 28 °C)—manage 

with shade so the air temperature does not exceed 91 °F 

(33 °C) maximum. Optimum temperatures of the rooting 

medium needs to be about 5 °F cooler [63 to 75 °F (17 to 24 

°C)] than optimum air temperatures.

Rooting Medium

A good rooting medium provides aeration and moisture, physically supports the cuttings, and promotes the 

development of fibrous root systems. A pH of 5.5 to 6.5 

is optimum for most plants, but acid-loving plants prefer 

4.0 to 5.0. Some common components of rooting media 

generally include a combination of two or more of the 

following: large-grade perlite, pumice, Sphagnum peat 

moss, sawdust, sand, coir, grit, or fine bark chips. The 

ideal rooting medium drains freely and does not become 

waterlogged from misting. Different rooting media components are used depending on the species being propagated. Selection of the rooting medium components 

influences rooting percentages and the quality of roots 

produced. Using very fine-grade or very coarse-grade 

sands tends to discourage the development of secondary 

roots. Roots that do form tend to be brittle and break off 

during transplanting. See Chapter 6, Growing Media, for 

more details on developing a good growing medium.

Mycorrhizal Fungi

Some growers inoculate the rooting medium with 

mycorrhizal fungi or other symbiotic organisms, which 

has improved rooting results with some species (Scagel and 

others 2003). See Chapter 13, Beneficial Microorganisms, 

for more information on mycorrhizae.

Nutrient Mist

Some difficult-to-root cuttings may remain in a special rooting environment for a long period of time. Over 

time, nutrients can be leached from the leaves by the 

long exposure to overhead misting, resulting in yellowing of the leaves or leaf and needle drop. In these cases, 

the application of a dilute, complete foliar fertilizer 

through the mist line can improve cutting vigor and may 

aid in rooting. Because some species respond favorably 

to nutrient mist while others are adversely affected, you 

will need to do some preliminary trials before treating all 

the cuttings. Also, be aware that excessive nutrients can 

encourage unwanted growth of mosses and algae on the 

medium surface.

Transplanting Cuttings From 

Special Rooting Environments

A few weeks after striking cuttings into the rooting 

environment, they should be inspected for root development. Using a trowel, carefully dig well below the end of 

the cutting and excavate it to examine for rooting. When 

cuttings have developed adequate root systems, they 

need to be hardened for transplanting outside the rooting environment. See Chapter 15, Hardening, for more 

information. The goal is to condition stem and leaf tissues and promote secondary root development before 

transplanting. Following these guidelines can harden 

cuttings:

• Gradually reduce the misting frequency over a period 

of 3 to 4 weeks.

• Increase the frequency and duration of ventilation in 

enclosed propagation systems.

• Do not let the rooting medium dry out completely.

After cuttings have hardened, transplant them into 

containers and transfer them to the nursery for additional 

growth (figure 10.19). Because cuttings are more expensive 

to produce than seedlings, it is important to handle them 

196 Tropical Nursery Manual
carefully at this stage. It is essential to avoid root damage 

by following these guidelines:

• Examine each cutting to ensure it has a root system 

capable of sustaining the cutting after transplanting. 

Cuttings with only a few slender roots or very short 

roots need to remain in the propagation bed for further root development (figure 10.20A).

• Transplant only on cool, overcast days or during early 

morning hours to avoid transplant shock.

• Transplant cuttings in an area of the nursery protected from wind and sunlight.

• Prepare containers, medium, labels, and transplanting 

tools before transplanting cuttings.

• Moisten the growing medium before transplanting to 

prevent tender roots from drying out.

• Remove cuttings from the rooting medium carefully 

and remove only a few at a time so roots will not dry 

out. Loosely wrap a moist paper towel around the root 

systems until they are transplanted.

• Handle cuttings carefully by holding the cutting by the 

stem and by leaving any rooting medium still attached to 

the root mass. Do not shake medium off the root system.

• Partially fill the container with moistened medium 

before inserting the cutting. Then add additional 

moistened medium and gently firm the medium with 

fingers without breaking the roots (figure 10.20B).

• Do not transplant the cuttings too deep or too shallow.

After transplanting the cuttings, they need to be 

placed in a shadehouse or protected from full sun and 

wind for at least 2 weeks. When the cuttings appear to 

be well established, gradually increase the level of sunlight by moving them to a different area of the nursery or 

by exchanging the shadecloth for one with a more open 

weave. After a couple of weeks, move sun-requiring species into full sun. Cuttings need to be closely monitored 

for any sign of stress. Adequate sunlight is needed for 

new shoot growth and adequate accumulation of carbohydrates before outplanting.

Other Methods of Vegetative 

Propagation

Besides stems and roots, several other portions of mother 

plants can be used to vegetatively propagate new daughters, 

and stems can be used in ways other than the traditional 

rooted cuttings.

Layering

Layering is a technique by which adventitious roots 

are formed on a stem while still attached to the plant. 

Vegetative Propagation 197

Figure 10.19—Cuttings can be transplanted into containers and 

moved to the nursery for hardening. Photo by Tara Luna.

Figure 10.20—Cuttings need to have 

enough developed roots that can support the cutting after it is lifted and 

transplanted outside the mist chamber 

or polypropagator (A). Cuttings with 

under-developed roots need to be left in 

the propagation bed longer to develop 

an adequate root mass (B). Photos by 

Tara Luna.

A B
Layering often occurs naturally without the assistance of 

a propagator. It is mostly used on those species that fail 

to root from stem or root cuttings. Four types of layering 

are commonly used in tropical nurseries: simple, French, 

air, and mound. In addition, drop and stack are two other 

layering techniques that could be used.

Simple Layering

Simple layering is used on species that produce many 

shoots annually. Long, low-growing flexible shoots are 

pegged down 6 to 9 in (15 to 23 cm) from the shoot tip, 

forming a “U” (figure 10.21A). The bottom of the U stem is 

girdled with a sharp knife and is covered with soil, sawdust, 

or other organic mulch, leaving the tip exposed. After a sufficient root system is formed, the new plant can be severed 

from the donor plant (figure 10.21B).

French Layering

French layering is similar to simple layering but uses 

a long, single branch that is pegged down to the soil surface (figure 10.22A). After a period of time, the pegs are 

removed and the branch is laid into a trench and buried 

up to the tips of the shoots with well-aerated soil and sawdust or mulch (figure 10.22B). After burying repeatedly, 

each shoot along the stem will form roots by the second 

year (figure 10.22C) and can be severed from the mother 

plant (figure 10.22 D).

198 Tropical Nursery Manual

Figure 10.22—French layering consists of pegging a long branch to the soil surface and allowing for new shoots to develop (A). 

After a period of time, the branch is buried in a deeper trench to encourage root development on each new shoot (B). Repeated 

burying (C) results in sufficient root development for plants to be severed from the mother plant (D). Graphic courtesy of Bruce 

McDonald and Timber Press, Inc.

Figure 10.21—In simple layering, the stem from a mother 

plant is pinned down and covered with soil or an organic mulch 

(A). New plants can be severed after there is sufficient shoot and 

root development (B). Graphic courtesy of Bruce McDonald and 

Timber Press, Inc.

B

A

B D

A C
Air Layering

Air layering is useful for producing a few plants of relatively large size. An advantage of air layering is that the 

rooted layer will be physiologically similar in age to the 

parent plant and will therefore flower and fruit sooner 

than a seedling or cutting. Air layering is used mostly on 

tropical fruit species, such as lychee, and to propagate 

rare and endangered tropical species. For optimum rooting, air layers are made on shoots produced during the 

previous season or during the mid- to late-active growing season on shoots from the current season’s growth 

(figure 10.23). For woody plants, stems of pencil-size diameter or larger are best. An area directly below a node is chosen, normally about 1 ft (30 cm) from the tip. Leaves and 

twigs on the stem are removed 3 to 4 in (7 to 10 cm) above 

and below this point. Air layering techniques differ slightly 

depending on whether the species is a monocot or a dicot.

The following steps describe air layering of monocots:

• Make an upward 1.0- to 1.5-in (2- to 4-cm) cut about 

one-third through the stem.

• Hold the cut open with a toothpick or wooden matchstick.

• Surround the wound with moist, unmilled Sphagnum

moss (about a handful) that has been soaked in water 

and squeezed to remove excess moisture.

• Wrap the moss with plastic and hold in place with 

twist ties or electrician’s tape. No moss should extend 

beyond the ends of the plastic.

• Fasten each end of the plastic securely, to retain moisture and to prevent water from entering. If it will be 

exposed to the sun, the plastic needs to be shaded.

The following steps describe air layering of dicots:

• With a sharp knife, make two parallel cuts about 1 in 

(2.5 cm) apart around the stem and through the bark 

and cambium layer.

• Connect the two parallel cuts with one long cut.

• Remove the ring of bark, leaving the inner woody 

tissue exposed.

• Scrape the newly bared ring to remove the cambial 

tissue to prevent a bridge of callus tissue from forming.

• Application of a root-promoting substance to the 

exposed wound is sometimes beneficial.

• Wrap and cover using the same procedure as that 

described for monocots.

Vegetative Propagation 199

Figure 10.23—After wounding the stem, an air layer is created by wrapping the area with peat moss (A), then enclosing this stem 

area in plastic wrap (B), and sealing the ends (C). After the layer has rooted (D), it can be severed from the stem and potted. Photos 

by Thomas D. Landis.

B

A C

D
For both monocots and dicots, the stem is severed below 

the medium after the rooting medium is filled with roots 

(figure 10.23D). The layer is then potted and needs to be 

kept shaded with adequate moisture until the root system 

becomes more developed.

Mound Layering

Mound layering or stooling involves selecting a 

young stock plant (figures 10.24A, 10.24B) and cutting 

back shoots to 2 to 4 in (5 to 10 cm) above ground level 

(figure 10.24C). Numerous shoots develop in consecutive growing seasons and are covered to one-half of their 

height with soil, sawdust, or other organic mulch (figure 

10.24D). This procedure is repeated three times as the 

shoots grow so that, by the end of the second or third 

growing season, the well-rooted shoots are unburied 

and are ready to plant as individuals (figures 10.24E, 

10.24F).

200 Tropical Nursery Manual

Figure 10.24—Mound layering or stooling involves selecting a young stock plant (A, B) and cutting back its shoots (C). After new 

shoots develop, they are covered to one-half of their height with soil, sawdust, or other organic mulch; this procedure is repeated 

three times (D), encouraging root development on the new shoots (E). After 2 or 3 growing seasons, the well-rooted shoots are ready 

to plant as individuals (F). Graphic courtesy of Bruce McDonald and Timber Press, Inc.

B

E

C

F

A

D
Drop Layering

Drop layering is very similar to mounding. Drop layering 

involves planting well-branched container plants deeply in 

the ground with only the tips of the branches exposed. New 

growth forms from the exposed branch tips, but the buried 

portions of the stems form roots along the stems.

Stacked Layering

Stacked layering is a new vegetative propagation 

method for rhizomatous species (Landis and others 2006). 

This technique takes advantage of the rapid and extensive 

root growth of seedlings and the fact that severed roots will 

form new shoots. In the spring, a stack of StyrofoamTM containers is created with a 1-gallon pot containing a seedling 

inserted in the top block. The lower StyrofoamTM containers are filled with growing medium with a thin layer of 

medium sandwiched between the blocks (figure 10.25A). 

After several months, the roots of the mother plant will 

have grown down through and colonized the cavities in 

the lower blocks. Running a sharp knife between the StyrofoamTM containers severs the roots, which then form new 

shoots (figure 10.25B). After a few months, the new plants 

can be transplanted into larger pots. Another set of filled 

StyrofoamTM containers can be situated below the block with 

the mother plant to start another propagation cycle. 

Tuberous Roots, Tubers, Rhizomes, and 

Crown Division

Tuberous roots, tubers, and rhizomes are specialized 

plant structures that function in the storage of food, 

nutrients, and water. Many culturally important tropical species, which are not easily grown from seeds, have 

these structures.

Tuberous roots are swollen secondary roots. Separating 

each tuberous root that has a section of the crown bearing 

a shoot bud produces new plants. Cassava, yams, and sweet 

potatoes, that are widely cultivated throughout Polynesia, 

produce tuberous roots.

Tubers are swollen modified stems that serve as underground storage organs. One well-known common tuber is 

the white potato. “Eyes” in white potatoes are actually nodes 

containing buds. Propagation by tubers involves planting 

the entire tuber or dividing it into sections containing at 

least one eye or bud. Pia, taro, wapato, and elephant taro 

are examples of tropical species that produce tubers.

Vegetative Propagation 201

Figure 10.25—Roots from the mother plant grow downward through the cavities of stacked StyrofoamTM containers (A). The 

roots are then severed (B), after which they develop new shoots (C). Adapted from Landis and others (2006).

A B C
Rhizomes are specialized stems in which the main axis 

of the plant grows horizontally or vertically at or below the 

soil surface. Many native tropical species, such as native 

begonias, reproduce by rhizomes and are easily propagated 

into larger numbers from a few nursery plants by divisions. 

Rhizomes vary in length and size according to species. Rhizomes are cut into sections with each containing at least 

one shoot bud or active shoot; some roots are attached to 

the bottoms of the rhizomes and are planted into containers individually. Rhizomes can also be planted in nursery 

beds and used as a source for bareroot stock for planting or 

for cultural uses such as basketry.

Crown division is an important method for propagating many native herbaceous perennials that produce multiple offshoots from the crown. Crown divisions are usually 

done just before active growth and flowering. Plants are 

dug up and cut into sections with a sharp knife, each with 

a substantial portion of the root system, and transplanted 

individually.

Stolons, Runners, and Offsets

Stolons, runners, and offsets are specialized plant 

structures that facilitate propagation by layering. Stolons 

are modified stems that grow horizontally above the soil 

line and produce a mass of stems. Runners are specialized 

stems that arise from the crown of the plant and grow horizontally along the ground and produce a plantlet at one of 

the nodes. Raised beds planted with species with stolons or 

runners can be an endless source of propagation material, 

and plants can be dug and potted individually or transplanted as bareroot stock.

Plants with rosettes often reproduce by forming new 

shoots, called offsets, at the base of the main stem or in 

the leaf axils. Offsets are cut close to the main stem of the 

plant with a sharp knife. If well rooted, an offset can be 

potted individually. Plantain and banana are often propagated with offsets (figure 10.26). Sever the new shoots 

from the mother plant after they have developed their 

own root systems. Nonrooted offsets of some species 

may be removed and placed in a rooting medium. Some 

of these offsets must be cut off, while others may simply 

be lifted from the parent stem.

Grafting and Budding

Grafting is the art of connecting two pieces of living 

plant tissues, the scion and the rootstock, together in such a 

manner that they will unite and grow as one plant. Grafting 

is used primarily in the tropics for mango, citrus, other 

tropical fruits, and forest tree seed orchards. In Hawai‘i, 

grafting has been used to propagate the highly endangered 

species, Kokia cookei (figure 10.27). It is also used to repair 

or to top-work existing trees to change varieties, as well as 

to produce new plants.

The scion is a short piece of shoot including several 

shoot buds, which, when united with the rootstock, forms 

the upper portion of the plant. The rootstock is the lower 

portion of the graft, which develops into the root system. 

If grafting is done high in the tree, the rootstock includes 

the roots, main trunk, and scaffold branches. The rootstock 

may be a seedling, rooted cutting, or an older tree.

Budding is grafting using a scion with only a single bud 

attached to a piece of bark. It may or may not include a thin 

sliver of wood under the bark. Budding is the most commonly used technique for propagating new plants, but it is 

also used to top-work existing trees to a new variety.

The rootstock and the scion must be compatible for 

successful grafting. Compatibility is never a problem when 

grafting within a clone. Grafting between clones within a 

202 Tropical Nursery Manual

Figure 10.26—Plantain and banana are often propagated with 

offsets. Photo by Ronald Overton.
species is usually successful. Grafting between species in a 

genus is sometimes successful and is most often seen in the 

genus Citrus. Grafting between genera within a plant family 

is rarely done and the chances of success are slim. Grafting 

between plant families is impossible for woody plants.

In addition to compatibility, a number of factors exist 

that contribute to a successful graft or bud union. Vascular 

cambium of the scion must be placed and held in intimate 

contact with the vascular cambium of the rootstock. The 

grafting operation must be done at a time when both the 

rootstock and the scion are in the correct physiological 

stage. Immediately after grafting, all cut surfaces must be 

protected from desiccation. Breaking of the union between 

the scion and the rootstock will usually manifest failed 

grafts. Grafts can fail because the rootstock and scion are 

not compatible, the rootstock and scion are not properly 

united, either the rootstock or the scion are not in the 

proper physiological state, or the grafted tissues dry out 

before the tissues grow together.

Some grafted species show excessive sap flow (bleeding) at certain times of the year. Excessive bleeding from 

the rootstock usually causes graft failure. Keeping recently 

performed grafts at 90 °F (32 °C) or lower with high humidity is essential. The graft union must also be protected from 

excessive drying or winds. In general, the more experienced 

the grafter, the better the success rate.

Grafting Tools

Sharp knives, sharpening stones, pruning shears, and 

saws are tools needed for grafting. These tools need to be 

kept very clean and only used for grafting. Knives need to 

be kept very sharp to minimize injury to the scion or rootstock. Dull knives strip and tear wood, leaving cuts that 

do not heal properly. Materials such as paraffin wax, budding rubber, and grafting tape can protect grafted tissues 

and seal cut surfaces of the graft. Aluminum foil wraps and 

plastic bags with twist ties are used in drier climates to protect the graft and provide a little extra humidity around the 

graft during union.

Collecting Scion Material

Budding is usually the preferred grafting method. For 

success, it is important to collect scion material with leaf 

buds and not flower buds. A small branch containing several buds suitable for grafting is called a bud stick. Bud 

sticks need to be collected when trees have well-developed 

buds. If buds have begun to swell or grow, the wood cannot be used successfully. Select parent trees of the desired 

variety that are disease free. Select straight, smooth bud 

sticks from 1-year-old wood that have 0.25- to 0.50-in (6- to 

13-mm) diameters and contain at least three buds or nodes. 

The best bud sticks usually come from the inside canopy of 

the tree. Seal about 6 mm of the end of each bud stick with 

melted wax or grafting paint. When the seal is dry, tie the 

bud sticks into small bundles and surround each bundle 

with moist paper towels or moist wood shavings to prevent 

desiccation. Label each bundle. The bundles and wrapping 

material can be kept inside plastic bags. Keep the bundles 

as cool as possible and do not allow them to dry out.

Types of Budding

Chip budding works well and can be done whenever 

mature buds are available. Chip budding is widely used 

for citrus propagation. The cuts on the scion and the rootstock must be exactly the same. The first cut on the scion 

Vegetative Propagation 203

Figure 10.27—Grafting has been used to propagate highly endangered species in Hawai‘i. Shown: Kokia cookei scion grafted onto the 

rootstock of Kokia drynarioides. Photo by Tara Luna.
Figure 10.28—Species like taro have been micropropagated 

to perpetuate certain clones and supply disease-free nursery 

stock. Photo by Tara Luna.

and the rootstock is made below a bud and downward at 

a 45-degree angle to a depth of about 0.12 in (3 mm). The 

second cut is started about the same distance above the bud 

and the knife is drawn downward to meet the first cut. If 

the bud scion happens to be narrower than the rootstock 

hole, line up one side of the bud scion tightly against the 

cut on the rootstock. The exact distances above and below 

the scion bud will depend on species. The entire graft needs 

to be wrapped with very thin (2-mil), clear polyethylene 

tape to prevent desiccation. If clear tape is unavailable, the 

graft can be wrapped with budding rubber and kept in a 

cool, shady location with high humidity. The key is to not 

let the bud dry out.

Slip budding is usually done during active growth. Like 

chip budding, mature buds must be available on the scion 

and the wood must have bark that will “slip.” Bark that 

“slips” will easily peel in one uniform layer, without tearing, from the underlying wood. The appropriate time to do 

this step depends on the species and local climate. The first 

cut on the rootstock is horizontal. The second cut, about 

twice as long as the first, is vertical, originating near the 

midpoint of the horizontal cut. Where the cuts meet, gently use the knife to slightly flare open flaps of bark. On the 

scion, detach the leaf below the bud but retain some of the 

petiole. Make the first cut about 0.5 in (12 mm) below the 

bud and draw the knife upward just under the bark to a 

point about 0.25 in (6 mm) above the bud. Grasp the petiole 

and make a second cut horizontally across the bud stick so 

that it intersects with the first cut. The bud and its accompanying wood, termed a bud shield, is then inserted under 

the “flaps” on the rootstock, and slid down to ensure that 

the scion makes intimate contact with the rootstock. Use 

a budding rubber to hold the stem, flaps, and bud shield 

firmly together. Do not cover the bud.

Finishing the Graft

Grafted surfaces must be held tightly in place using a 

budding rubber or grafting tape. This wrap must either 

breakdown by weathering (as budding rubbers do) or 

must be removed in 2 to 3 weeks after the union has 

healed. If the material does not break down and is not 

removed, it will girdle the rootstock. After the union has 

healed, the portion of the rootstock above the graft must 

be cut away to force the scion bud to grow. Remove any 

unwanted sprouts as soon as they are visible. Unwanted 

sprouts can be easily rubbed off with fingers.

Micropropagation

Micropropagation is a process used to propagate plants 

using very specialized tissue culture techniques. Tissue culture is the procedure for maintaining and growing plant tissues and organs in an aseptic culture in which the environment, nutrient, and hormone levels are tightly controlled. 

A small piece of vegetative material called the explant is 

used to create a new, entire plant. Rare or greatly endangered tropical native species have been micropropagated to 

increase the number of individuals for restoration projects 

when other methods of propagation have been limiting or 

failed. Micropropagation has also been used as a method 

to offer plants in the nursery trade in order to preserve 

them from poaching and eventual extirpation from wild 

populations (figure 10.28). Micropropagation works well 

for some species and poorly for others. For some native 

plants, such as orchids, it is one of the only options for 

successful propagation. Most native plant nurseries do 

not have an elaborate tissue culture facility because of the 

high cost, although small-scale micropropagation can be 

done with minimal equipment in a clean room.

204 Tropical Nursery Manual
References

Landis, T.D.; Dreesen, D.R.; Dumroese, R.K. 2003. Sex and the 

single Salix: considerations for riparian restoration. Native 

Plants Journal. 4: 111–117.

Landis, T.D.; Dreesen, D.R.; Pinto, J.R.; Dumroese, R.K. 2006. 

Propagating native Salicaceae for riparian restoration on the 

Hopi Reservation in Arizona. Native Plants Journal. 7: 52–60.

Landis, T.D.; Tinus, R.W.; Barnett, J.P. 1999. The container tree 

nursery manual: volume 6, seedling propagation. Agriculture 

Handbook 674. Washington, DC: U.S. Department of Agriculture, Forest Service. 167 p.

Longman, K.A. 1993. Rooting cuttings of tropical trees: propagation and planting manuals, volume 1. London, United Kingdom: 

Commonwealth Science Council. 137 p.

Scagel, C.F.; Reddy, K.; Armstrong, J.M. 2003. Mycorrhizal fungi 

in rooting substrate influences the quantity and quality of roots 

on stem cuttings of hick’s yew. HortTechnology. 13: 62–66.

Vegetative Propagation 205
206
207

Water is the single most important biological factor affecting plant growth and 

health. Water is essential for nearly every plant process: photosynthesis, nutrient 

transport, and cell expansion and development. In fact, 80 to 90 percent of a seedling’s weight is made up of water. Therefore, irrigation management is the most critical aspect of nursery operations (figure 11.1). In tropical nurseries, managing water 

depends on local climate. Finding and storing high-quality water is a challenge in dry 

climates, whereas controlling high humidity can be a problem in the humid tropics.

Determining how, when, and how much to irrigate is a crucial part of nursery 

planning and day-to-day operations. One missed irrigation can cause serious injury 

and even death to plants, especially during the establishment phase. Adequate 

watering is particularly important with container plants whose roots cannot access 

water beyond their container and therefore are entirely dependent on receiving 

enough water through irrigation. Excessive watering and humidity are also problematic; they are a major cause of root diseases and contribute to other problems 

with seedling growth.

Tropical nurseries typically grow a wide range of species with different water 

requirements. In addition, the distinct plant growth phases (establishment, rapid 

growth, and hardening) require different watering regimes. Designing an effective 

and efficient irrigation system is based on which types of irrigation practices and 

systems best serve plant needs at each developmental phase. The nursery might have 

various propagation areas and corresponding irrigation zones that provide for the 

changing needs of plants during all phases of growth. For example, the same nursery 

might have a mist chamber for the germination phase, a subirrigation system in the 

growing area, a selection of rare plants that receive daily hand watering, and some 

large plants under drip lines. The best design for any irrigation system will come 

from understanding the needs of the plants, the factors that affect water availability, 

and the details of how, when, and why to water.

Water Quality and Irrigation

Thomas D. Landis and Kim M. Wilkinson 11

Facing Page: Water is a precious resource and the most important biological factor for plant health and growth. Pe‘epe‘e Falls, 

Wailuku River State Park, Hawai‘i. Photo by Douglass F. Jacobs. 
Sources of Irrigation Water

Tropical plant nurseries can use water from several 

different sources, including rivers, ponds or reservoirs, 

rainwater, groundwater, and municipal sources. New 

nurseries need to evaluate the quantity, quality, and seasonal availability of all potential water sources. Surface 

water, such as rivers and streams, are typically dammed 

or diverted into ponds that have enough capacity to meet 

the water demands of the nursery. Groundwater can be 

pumped for nursery use, but generally the water is stored 

in a pond or reservoir. For either surface or groundwater, 

a hydrologic survey and analysis of local water rights needs 

to be conducted before nursery development. Surface water 

sources that have flowed through agricultural land need to 

be tested for waterborne pests or herbicides and may need 

to be treated. A wide variety of nursery pests have been 

detected in water supplies including fungi, nematodes, 

viruses, and bacteria (Zeng and others 2009). The water 

mold fungi (Pythium and Phytophthora species) are particularly destructive because they have a mobile stage.

Municipal water can be used for nursery purposes, but it 

may have been chemically treated with chlorine or fluoride. 

Low levels of chlorination guard against human pathogens, 

and studies have shown water with 2 ppm of free chlorine will 

not injure woody species (Cayanan and others 2008). Fluoride is added to some municipal drinking water at around 1 

ppm to prevent cavities in children (Fawell and others 2006). 

Fluoride phytotoxicity has been documented for some nursery plants. Leaf damage to ti (Cordyline fruticosa) plants has 

been reported in a Hawaiian nursery, and the nursery recommends that irrigation water contain less than 0.25 ppm 

fluoride (Kaapuni Nursery 2011).

Rainwater is an attractive source of high-quality water 

for tropical nurseries if enough can be stored to supply all 

the needs. In high-rainfall areas, water can be collected 

from the roofs of buildings and then stored in tanks until 

needed. A special rainwater collection system is being used 

to supply water for irrigation at the native plant nursery 

at Volcanoes National Park in Hawai‘i (figure 11.2A). In 

these ground-level systems, the water must be pumped to 

generate enough pressure to run sprinklers in the nursery. 

In mountainous regions, rain water can be collected and 

stored in ponds or tanks above the nursery. Enough water 

pressure can be generated by the difference in elevation 

to run small microsprinklers or drip irrigation systems in 

a nursery. The water pressure can be calculated (figures 

11.2B, 11.2C) or measured directly at the point of use in the 

nursery. A full discussion of all types of irrigation designs 

and calculations is available in Stetson and Mecham (2011).

208 Tropical Nursery Manual

Figure 11.1—A supply of good-quality water is one of the 

most critical requirements for a native plant nursery. Photo by 

Brian F. Daley.

Advantages of a Well-Designed 

Irrigation System

• Better plant quality and health

• Lower labor costs

• Improved crop uniformity and reliability

• Reduced runoff and waste of water

Attributes of Good Water Management

• Reliable source of water

• Efficient use of water

• Flexible approach tailored to the changing needs 

of the species grown and their development 

phases

• Responsible reuse, recycling, and management of 

any runoff water
Water Quality

Irrigation water quality is a critical factor in initial site 

selection and affects all phases of nursery management. 

Improving poor-quality irrigation water is expensive, often 

prohibitively so. Therefore, water quality needs to be a 

primary consideration during nursery site evaluation. 

For irrigation purposes, water quality is determined 

by two factors:

• The types and concentrations of dissolved salts (total 

salinity and individual toxic ions).

• The presence of pests (pathogenic fungi, weed seeds, 

algae, and possible pesticide contamination).

Salts

For our purposes, a salt can be defined as a chemical 

compound that releases charged particles called ions when 

dissolved in water. Some salts are fertilizers that increase 

plant growth rates, while other salts can reduce growth or 

even cause injury or death. For example, sodium chloride 

(ordinary table salt) dissolves into harmful ions that can 

damage or even kill plant tissue.

An excess of dissolved salts in nursery irrigation water 

can clog nozzles (figure 11.3A), accumulate in growing 

media, and eventually harm plant tissue (figure 11.3B), 

The most characteristic symptom of high salinity is burn or 

scorch of leaf margins or tips (figure 11.3C). Symptoms can 

Water Quality and Irrigation 209

Figure 11.2—Rain collection systems are possible in high-rainfall climates (A). If the storage tanks are located at higher elevations, enough static pressure can be generated (B) to operate drip or microsprinkler systems in the nursery (C). Photo A by Thomas 

D. Landis, and illustrations B and C modified from Stetson and Mecham (2011) by Jim Marin. [Metric conversions: 1 in = 2.5 cm; 

1 ft = 0.91 m; 1 psi = 0.007 MPa; 1 pound = 0.45 kg.]

B

C

A
vary with species but can include foliar tip burn, scorching or bluish color on leaves, stunting, patchy growth, and 

eventual mortality. The principal damage of high salinity is 

reduced growth rate, which usually develops before more 

visible symptoms become evident. 

Other ways excess dissolved salts may affect nursery 

crops include the following—

• Water availability to the plant is reduced resulting 

in growth loss.

• Some ions (sodium, chloride, boron, and fluoride) 

are directly toxic to plants.

• Other ions (calcium) affect mineral nutrient 

availability.

• Other ions (bicarbonate or iron) cause salt crusts 

or staining.

An excess of dissolved salts in the water can be the 

result of a number of factors. The local climatic or geologic 

influences at the nursery’s location are significant. In arid 

or semiarid climates where evapotranspiration exceeds 

precipitation, salts naturally accumulate in the soil, and 

ground water irrigation sources are often high in salt content. In coastal areas, saltwater intrusion can lower water 

quality. On Guam, the geology of the northern half of the 

island is formed from uplifted coral whereas the southern 

half is volcanic (Gingerich 2003). From a nursery standpoint, 

this distinction is that the groundwater in the north is very 

high in pH and calcium and in the south, the water is low in 

pH and dissolved salts. High fertilization rates or poor irrigation practices can also lead to salinity problems. Studies 

have shown that soluble salt levels double when the growing 

medium dries from 50- to 25-percent moisture content.

It is expensive to remove salts from irrigation water, so 

ideally the nursery needs to be established on a site where 

water salinity is within acceptable levels. Test results for 

salinity are traditionally expressed as electrical conductivity (EC); the higher the salt concentration, the higher the 

EC (table 11.1). The EC can be checked at the nursery using 

a conductivity meter or by sending water samples to a local 

laboratory. The most commonly used units in irrigation 

water quality are micromhos per centimeter (abbreviated 

as μmho/cm and pronounced “micro-mows”) and the 

International System of Units of microsiemens per centimeter, which are equivalent. Microsiemens per centimeter 

(abbreviated as μS/cm) will be used as the standard EC unit 

in this handbook. General guidelines for salinity ranges are 

in table 11.1.

Irrigation water salinity tests need to be conducted before 

nursery establishment and retested periodically thereafter. 

210 Tropical Nursery Manual

Figure 11.3—Agricultural water quality is determined by the level of soluble salts because they can build up on irrigation nozzles 

(A), accumulate in containers, usually around the drainage holes (B), and eventually “burn” seedling foliage (C). Photos by Thomas 

D. Landis.

Quality index Optimal Unacceptable

pH 5.5 to 6.5

Salinity (μS/cm) 0 to 500 >1,500

Sodium (ppm) <50 >50

Chloride (ppm) <70 >70

Boron (ppm) <0.75 >0.75

Fluoride (ppm) <1.00 >1.00**

Iron (ppm) <1.00 >1.00

Table 11.1—Water-quality standards for nursery irrigation 

water. Adapted from Landis and others (1989) and Robbins 2011.

** Sensitive species may be damaged at lower levels

A B C
EC tests were conducted at native plant nurseries in some 

of the Micronesian islands and the results reflect local geology (table 11.2). As previously mentioned, the northern 

part of Guam is limestone and the EC is much higher than 

the volcanic soils in the southern part. The water in Saipan 

has very high EC readings because of limestone soils, and 

this marginal water quality can be seen in the nursery.

Water tests are particularly important in areas with high 

salinity because the addition of fertilizer could raise salinity 

to unacceptable levels (figure 11.4). In these cases, a nursery 

would need to be careful to use very dilute fertilizers, lownutrient concentrations of organic fertilizers, or controlledrelease fertilizers to keep salinity within acceptable ranges. 

See Chapter 12, Plant Nutrition and Fertilization, for details. 

Horticultural practices such as increasing the porosity of the 

growing medium and leaching more frequently when watering can also help alleviate the effects of saline water.

Pests

Tropical nurseries that use irrigation water from surface 

water sources such as ponds, lakes, or rivers may encounter 

problems with biotic pests; that is, weeds, pathogenic fungi, 

moss, algae, or liverworts. Surface water that originates from 

other nurseries or farmland is particularly likely to be contaminated with water-mold fungi, such as Pythium and Phytophthora, which cause damping-off. Recycled nursery irrigation water should also be suspect and needs to be analyzed. 

Many weed seeds and moss and algal spores are small enough 

to pass through the irrigation system and can cause real problems in nurseries. Chlorination and some specialized filtration systems may remove many disease and pest organisms 

from irrigation water, as discussed in the following section.

Irrigation water, especially in agricultural areas, may be 

contaminated with residual pesticides. Herbicides applied 

to adjacent cropland or to control aquatic weeds in reservoirs can affect ground water. Potential sources of irrigation water need to be tested for pesticide contamination 

when a nursery site is being evaluated.

Testing Water Quality

A water-quality test needs to be done before the nursery site selection is finalized to ensure adequate quality 

and quantity of water will be available before the nursery 

is established. A water-quality test can be repeated when 

the nursery is established and again at yearly intervals. A 

complete analysis of irrigation water quality consists of a 

salinity evaluation listing the concentrations of sodium, 

chloride, and boron, which are reported in parts per million (ppm). In addition to the individual ion concentrations, the analysis needs to include three standard waterquality indices: EC, toxic ion concentrations, and pH. 

Most labs have a standard irrigation water-quality test, 

which will cover all these concerns.

Water Quality and Irrigation 211

Island Electrical conductivity 

(μS/cm)

Quality rating

(see table 11.1)

Guam (North) 840 Marginal

Guam (South) 150 Good

Saipan 1,860 Poor

Yap 185 Good

Palau 60 Good

Pohnpei 

(domestic) 125 Good

Pohnpei 

(rainwater) 16 Good

Table 11.2—Water-quality test results from Micronesian Islands.

Figure 11.4—When soluble fertilizers are injected into the irrigation system, salinity levels are cumulative. For example, a 

nursery with a base irrigation salinity of 500 μS/cm has good 

quality, but after soluble fertilizer is added, the total salinity 

can reach into the zone of caution. Illustration from Dumroese 

and others (2008).

μS/cm
Irrigation water also needs to be tested for the presence of 

pathogenic fungi during the site selection process, and again 

if a problem is observed at a later date. Most plant pathology 

laboratories can conduct bioassays of irrigation water.

Testing for residual herbicides is also possible but can be 

expensive because of the sophisticated analytical procedures 

required. Because of the different chemical structures of various pesticides, a separate analysis for each suspected pesticide is usually required. Therefore, pesticide tests are generally considered only when a specific problem is suspected.

To collect an irrigation water sample, use a clean plastic 

bottle with a firm, watertight lid. A 16-fluid-ounce (475-

ml) container is ideal for most water tests. To begin, let the 

water run for several minutes (figure 11.5), and then rinse 

the sample bottle well before collecting the sample. Label 

the sample bottle properly with a waterproof marker before 

sending it to the analytical laboratory. The sample needs 

to be sent away for testing as quickly as possible but can 

be stored under refrigeration for short periods, if necessary. Most laboratories charge $25 to $50 and will provide 

results within a few weeks.

Water Treatments

Establishing the nursery on a site with tested, goodquality water is the best way to preclude water-related 

problems. If existing water quality is poor, methods such 

as deionization and reverse osmosis can treat and improve 

irrigation water, but they are often prohibitively expensive 

and not feasible for most tropical plant nurseries. To correct or safeguard against minor problems with otherwise 

good-quality water, however, some treatments are low cost 

and highly effective for container nurseries.

Chlorination

Chlorination can be used to kill fungi, bacteria, algae, 

or liverworts introduced through the irrigation system (Zheng and others 2009). Studies have shown that 

Pythium and Phytophthora can be controlled with 2 ppm 

of free chlorine without injury to woody species (Cayanan 

and others 2008). 

Filtration

Filtration is used to remove suspended or colloidal 

particles such as very fine sand or silt. Filtration prevents 

problems such as plugging or damaging irrigation or fertilization equipment. Filters also remove unwanted pests 

such as weed seeds or algae spores. Two types of filters 

are commonly used in nurseries: granular medium filters 

consist of beds of granular particles that trap suspended 

material in the pores between the particles and surface filters use a porous screen or mesh to strain the suspended 

material from the irrigation water. Granular medium filters can be used to remove fine sand or organic matter 

and are constructed so that they can be backflushed for 

cleaning. Surface filters include screens or cartridges of 

various mesh sizes to remove suspended material (figure 

11.6A); screens must be physically removed and cleaned 

whereas cartridge filters are not reusable and must be 

regularly replaced. Jones (1983) recommends cartridge 

filters because they are easy to change. Backflushing 

screens or granular medium filters is not practical with 

many nursery irrigation systems.

Filters need to be installed at a location before the 

water passes through the nutrient injector to intercept 

sand particles that can cause excessive wear or plug 

valves (figure 11.6B). Handreck and Black (1984) recommend using filters small enough to remove particles 

212 Tropical Nursery Manual

Figure 11.5—Water quality needs to be tested before nursery 

establishment and then every few years to make certain that the 

quality has not changed. Photo by Brian F. Daley.
greater than 5 microns in diameter, which will take care 

of most suspended materials (figure 11.6C). Specialized 

filtration systems, such as those manufactured by Millipore Corporation, can remove particles around 1 micron 

in diameter; such a system is therefore capable of removing 

some disease organisms and most suspended solids. More 

sophisticated filtration systems are relatively expensive and 

require frequent maintenance (Jones 1983).

Water Quantity

The amount of water necessary to produce a crop of 

container plants depends on many factors, such as climate, type of growing structures, type of irrigation system, growing medium, plant species, number of plants 

being grown, and container size. The amount of water to 

grow a crop will vary tremendously between humid and 

arid locations. Some examples of water use data from 

nurseries in the mainland United States are provided in 

table 11.3.

Remember that a nursery also needs water for operational requirements other than irrigating crops. Mixing 

growing media, cleaning containers, structures, and equipment, and providing for staff personal water needs all 

increase water use. Also, a nursery that starts small may 

choose to expand. Therefore, ensure that an abundance of 

water is available to meet present and future needs.

Even in cases in which the nursery has access to a 

steady, reliable, high-quality water source, a backup system is always a good idea in case of emergency. A prudent 

investment is a backup water storage tank containing 

sufficient water to meet the nursery’s needs for at least 1 

week (figure 11.7). Backup systems may be pumped into 

the normal irrigation system, but it is advantageous to 

locate the storage tank upslope so that water can be supplied by gravity in case of power failure.

Water Quality and Irrigation 213

Figure 11.6—Cartridge filters (A) are an effective and inexpensive way to treat irrigation water: they need to be installed before 

the fertilizer injector (B) and can remove sand, silt, and fungal and 

algal spores (C). Illustration A from Dumroese and others (2008), 

photo B by Thomas D. Landis, and illustration C adapted from 

Tchobanoglous and Schroeder (1985) by Jim Marin.

Chlorination with Household Bleach To 

Treat Water for Pests

• Mix household bleach (5.25-percent sodium 

hypochlorite) at a rate of 2.4 ounces of bleach per 

1,000 gal of water (18 ml per 1,000 L).

• This low dose (about 1 ppm chlorine) was not 

found to be phytotoxic to a wide range of plant 

species, and it has been successful in controlling 

moss and liverwort on noncrop surfaces.

B

C

A
erable variation in growing medium water content from one 

container to another. This distribution problem becomes 

even more critical when the plants become larger and their 

foliage begins to intercept water before it can reach the surface of the container. Subirrigation (described later in this 

chapter) and hand watering are sometimes used to overcome 

this problem. Because small containers have a correspondingly small volume of growing medium, they have limited 

moisture reserves and require frequent irrigation, especially 

in times of high evapotranspirational losses.

Determining How Much 

To Irrigate

When irrigating container nursery crops, it is important to apply enough water during each event to more 

than saturate the medium so that a small amount of 

leaching occurs. In other words, apply enough water so 

Factors Affecting Water 

Availability to Plants

Plant water use is affected by environmental conditions such as humidity, temperature, season, and the 

amount of sunlight the plants receive. The growth phase 

of the crop will also affect the rate of evaporation and 

transpiration. During seedling germination and early 

emergence, evaporation is the primary cause of water loss 

(figure 11.8A). After the seedling’s roots occupy the container, however, transpiration becomes the primary force 

for water loss (figure 11.8B).

The type of growing medium also influences water availability and use. Common components of artificial media 

behave very differently than soil. Peat moss and vermiculite have a high water-holding capacity, whereas perlite and 

pumice do not. Water infiltration and drainage are much 

higher with artificial media than with mineral soils. The 

average pore size of the growing medium is the most significant influence. All things being equal, a finer textured 

growing medium with a smaller average pore size holds 

more water than a coarser textured medium does (figure 

11.9). See Chapter 6, Growing Media, for more details on 

this topic.

Container type, volume (table 11.3), and shape also affect 

water availability. Water in a container behaves differently 

than water in unconfined soil because it does not drain completely, resulting in a layer of saturated medium at the bottom (figure 11.8B). The height of this saturated medium is a 

function of the growing medium, but taller containers will 

have a smaller proportion of saturated medium than shorter 

ones (see Figure 7.2 in Chapter 7, Containers).

The small top opening of some containers in relation to 

their volume can make it difficult to get water into the containers with typical sprinkler irrigation, resulting in consid214 Tropical Nursery Manual

Nursery and location Container type and volume

Irrigation water use per week (gallons [liters])

Establishment phase Rapid growth phase

University of Idaho, Moscow Ray Leach Cone-tainerTM

4 in3 (66 ml) 10 (38) 15 (57)

Mt. Sopris, Colorado Ray Leach Cone-tainerTM

10 in3 (172 ml) 15 (57) 50 (189)

University of Idaho, Moscow StyroblockTM 

20 in3 (340 ml) 60 (227) 125 (473)

Table 11.3—Typical irrigation use in forest and conservation nurseries for a crop of 1,000 conifer seedlings on the mainland of the 

Western United States. Adapted from Dumroese and others (2008).

Figure 11.7—A backup water storage tank containing sufficient water to meet the nursery’s needs for at least 1 week is a 

prudent investment. Photo by Thomas D. Landis.
that some drips out the bottom of the container (figure 

11.10A), but not so much that water streams out the bottom. If the irrigation period is too short, the water will 

not reach the bottom of the container and a layer of dry 

growing medium underneath will exist (figure 11.10B). 

If the growing medium is not completely saturated after 

irrigation, the seedling will not develop roots in the dry 

medium at the bottom of the container, resulting in a poorly 

formed plug. In addition, fertilizer salts will accumulate in 

the medium and cause salinity damage or “fertilizer burn.”

The general rule for sprinkler, overhead, hand-watering, or microirrigation is to apply approximately 10 percent more water than is needed to completely saturate the 

entire growing medium profile during irrigation. The best 

procedure to ensure that some water is dripping out of 

the bottom of the container during or immediately after 

irrigation is by direct inspection.

The amount of irrigation to apply varies during the growing season because of the stages of seedling development 

and the horticultural objectives of the nursery manager. As 

discussed in Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records, plants go through three phases 

of development: establishment, rapid growth, and hardening. Irrigation is an important strategy for managing plant 

growth and health as the crop moves through these phases.

Irrigating During the 

Establishment Phase

Immediately after the sown containers are placed in 

the growing area, the growing medium needs to be completely saturated. Thereafter, watering needs during establishment should be monitored carefully and tailored to the 

Water Quality and Irrigation 215

Figure 11.9—Water is held in the pores between particles by 

capillarity. For any growing medium, the larger the particles, the 

less water will be held. Plants can only use capillary water; gravitational water drains away, and hydroscopic water is held too tightly 

for plant use. Illustration from Dumroese and others (2008).

Figure 11.8—The amount and type of water use changes dramatically during the growing season. During the establishment 

phase, a relatively small amount of water is used by evaporation 

(A) but, when the seedling fully occupies the container during the 

rapid growth phase, a much greater amount is used for transpiration with relatively little lost to evaporation (B). Illustration 

adapted from Landis and others (1989).

A B

Figure 11.10—Apply enough water during irrigation to fully saturate the growing medium profile and permit some leaching out 

the bottom of the container. Some water will drip out of the bottom of each container if the plants were watered sufficiently (A). 

Insufficient irrigation will result in a dry layer within the growing 

medium (B). Photos by Thomas D. Landis.

A B
needs of the species. Some species require nearly continuous misting (figure 11.11) and may even benefit from a fog 

chamber. Other species require less water, but, nevertheless, inadequate or too-infrequent irrigations will cause the 

seeds to dry out, which will decrease germination success 

or even cause total crop loss. On the other hand, excessive 

irrigation may create overly wet conditions, promoting 

damping-off and decreasing germination.

To help determine the watering regime for a given species, consider the water availability in the plant’s native 

habitat (such as rainforest, cloudforest, or dryland) at the 

time of year when this species usually germinates. Remember, until the seeds germinate and begin to grow, water loss 

is primarily from evaporation from the top of the container. 

Irrigation during this period, therefore, must be applied with 

the goal of replenishing the moisture in the thin surface layer 

of the medium. This practice is usually best accomplished 

by periodic misting or light irrigation with a very fine spray 

nozzle, which also protects germinating seeds from being 

moved or damaged by the force of the water. In some cases, 

if seeds are mulched, irrigation may not be necessary for the 

first week or so of the germination phase. Scratch through 

the surface of the mulch or grit to check the water status of 

the medium and ensure the medium is moist enough.

Irrigation can also be used to control the temperature 

around germinating seeds. High temperatures on the surface 

of the growing medium can injure germinants, particularly 

those covered by dark-colored mulches. If temperatures 

exceed 86 °F (30 °C), light misting will help keep germinants 

cool through evaporative cooling. This practice is sometimes 

called “water shading.” When misting for cooling, do not 

add too much water to the medium, only enough to dissipate 

heat around the seedling. After plants develop thicker stems, 

they become more resistant to heat injury. Do not use water 

shading in nurseries with saline water because salts can build 

up in surface layers of the growing medium. 

Irrigating During the 

Rapid-Growth Phase

After the seedling’s roots have expanded throughout the 

container, the amount of water lost through transpiration 

increases greatly and so irrigations must be longer and more 

frequent. As seen in table 11.3, water usage can double or 

even triple during the rapid-growth phase. Each plant needs 

to be watered thoroughly (until some water drips out the 

bottom). Regular saturation is best for some species whereas 

others might benefit from periods of slight moisture stress 

between watering. No plants should ever be allowed to dry 

out completely (figure 11.12). Nursery managers need to be 

aware of the varying water requirements for different species 

and adjust irrigation practices accordingly. Grouping species 

together by their water requirements (“wet,” “moderate,” or 

“dry”) makes this practice much easier.

During the rapid growth phase, the leaves of plants begin 

to form a tight canopy that causes a significant reduction in 

the amount of irrigation that can reach the growing medium 

surface if delivered from above. This “umbrella effect” is particularly serious with broadleaved species. Water will often 

drip through the foliage irregularly so that one container is 

fully saturated whereas the one right next to it may receive 

216 Tropical Nursery Manual

The key to irrigating during the 

establishment phase: keep the growing 

medium “moist, but not saturated.”

Figure 11.11—During the establishment phase, watering 

needs to be tailored to the requirements of each species. For 

many species, mist nozzles in a special germination chamber 

generate a fine spray, which provides enough water for germination and also protects young germinants from heat injury. 

Overwatering must be avoided to prevent disease problems. 

Photo by Brian F. Daley.

Water Use Considerations for Plants 

Grown in Containers

• Root volume is proportional to plant size.

• Plants in containers do not have the ability to 

“forage” for water outside the root zone of the 

container.

• Because containers are relatively small, moisture 

reserves can be used up quickly in hot weather.

• Containers have a small top opening relative to 

volume that makes it challenging to apply equal 

amounts of water per container.
very little water. To compensate for foliage interception, 

growers tend to irrigate more frequently and longer. This 

practice still results in uneven irrigation and wasted water, 

because the water runs off the leaves and onto the floor. The 

types of irrigation systems discussed later in this chapter will 

help address this issue for broadleaved species, particularly 

through subirrigation and hand watering practices.

The rapid growth phase is also the time when liquid 

fertilizers are most concentrated and water loss through 

transpiration is high, so growers must monitor for salt 

accumulation.

Irrigating During the Hardening Phase

Manipulating irrigation frequency is an effective way to 

initiate the hardening of plants before shipment and outplanting. Because seedling growth is so critically tied to 

moisture stress levels, growers can slow shoot growth and 

increase general resistance to stress by inducing moderate 

water stress. This “drought stressing” procedure consists 

of withholding irrigation for short periods of time until the 

plants can be seen to wilt or until some predetermined moisture stress is reached. After this stress treatment, the crop is 

returned to a maintenance irrigation schedule.

Implementing moisture stress, however, can be challenging with tropical plants because (1) hardiness is affected by 

other environmental conditions, (2) considerable variation 

in growing medium moisture content can exist among containers, and (3) if the growing medium is allowed to dry too 

far, it can become hydrophobic and difficult to rewet.

Water stressing must be done correctly and conscientiously, and there is no substitute for experience. Most of 

the water stress research has been done with commercial 

conifers, and good guidelines have been published for 

monitoring container weights (for example, Landis and 

others 1989). It is unfortunate that little is known about 

the response of most tropical native plants. Inducing moisture stress, therefore, can be risky if the plant’s tolerance is 

unknown. Drought stressing simply does not work for some 

species and in some environments. Growers need to conduct 

their own trials of operational moisture stressing to determine the effect on their own species in their respective 

growing environments. Careful scheduling and communication with other nursery workers is essential (figure 

11.13). Be sure to keep detailed records of how the crops 

respond.

In spite of these caveats, the induction of mild moisture 

stresses should be considered as a horticultural technique 

to manipulate seedling physiology and morphology. A further discussion of the hardening process, including moisture 

stress, is provided in Chapter 15, Hardening.

Fertigation

Irrigation is critical to the proper application of fertilizers, especially when injecting liquid fertilizer solution into 

the irrigation system—a practice called “fertigation.” Fertigation can be used with many different types of irrigation 

systems, from hand watering to automated sprinkler or drip 

systems. Fertilizer injectors range from simple, low-cost 

siphons for hand watering to sophisticated pumps for automated sprinklers. Because it can be designed to apply the 

proper mineral nutrients at the proper concentration and at 

the proper time, fertigation has several advantages over other 

types of fertilization. Remember that fertilizers are salts and 

that injecting liquid fertilizers adds to the base salinity level 

of the irrigation water. Frequent leaching with regular irrigation water (“clearwater flush”) is needed to push excess 

salts out the bottom of the container. One of the first signs 

of a salinity problem is salt crust around the drainage holes 

(figure 11.3B). See Chapter 12, Plant Nutrition and Fertilization, for more information on fertigation and how it can 

be applied in tropical native plant nurseries.

Determining When to Irrigate

It is absolutely necessary to regularly monitor the moisture status of growing media. In small containers, the limited 

volume of moisture reserves means that critical moisture 

stresses can develop quickly.

Visual and tactile assessments are the most common 

method of monitoring irrigation effectiveness. Monitoring 

can also include formal or informal assessments of container 

weight. In addition, various tools such as tensiometers, 

electrometric instruments, balances, commercial moisture 

meters (figure 11.14), or pressure chambers can be used to 

Water Quality and Irrigation 217

Figure 11.12—These seedlings are suffering from severe water 

stress because of improper irrigation. Seedlings use a lot of water 

during the rapid growth phase and frequent irrigation is needed to 

prevent growth loss. Photo by Thomas D. Landis.
monitor irrigation efficacy (Landis and others 1989). Any 

equipment-based method must also be supported by actual 

observation (visual and tactile) and the grower’s experience.

Visual and Tactile Assessment of 

Growing Medium Moisture

Most nurseries successfully monitor irrigation efficacy 

based on the feel and appearance of the plants and the growing medium (figure 11.15A). The best technique is to observe 

the relative ease with which water can be squeezed from the 

medium and attempt to correlate this moisture condition 

with plant appearance and container weight (figure 11.15B). 

This process requires a lot of experience and is very subjective but can be very effective when used by a knowledgeable, 

experienced nursery manager.

Looking at the root systems or the growing media may 

involve damage to the plants that are examined, especially if 

they must be pulled from their containers. This practice may 

be necessary during the learning phase of growing a new crop. 

With time and experience, nondestructive indicators such as 

the appearance of the plant, the look and feel of the growing 

medium, and the weight of the containers will be practiced 

most of the time, and the need for destructive sampling will 

be reduced or eliminated.

Monitoring Container Weights

Monitoring container weight is one of the few objective, 

nondestructive, and repeatable techniques for monitoring irrigation in container nurseries. Container weight is 

also the best way to determine irrigation needs early in the 

218 Tropical Nursery Manual

Figure 11.13—Because so little is known about the response 

of most tropical plants to water stress, good scheduling, recordkeeping, and communication are critical to determine plant 

needs. Photo by Thomas D. Landis.

Figure 11.14—Many types of tools can be used to help test and 

assess the effectiveness of water application, including this moisture meter. Visual and tactile assessments are more prevalent in 

tropical plant nurseries, however, and can be quite effective when 

carried out by trained staff. Photo by Thomas D. Landis.
Water Quality and Irrigation 219

Figure 11.15—Growers can monitor the need for irrigation by careful observation of plant condition (A) so that water can be applied 

before plants are seriously wilted (B). Feeling the weight of containers can also be an effective way to determine when to irrigate (C). 

Monitoring container weights is a standard method to assess irrigation needs in container crops (D). Photos A, B, and C by Tara Luna, 

and photo D by Thomas D. Landis.

B

C

A

D
growing season before plants are large enough to show 

moisture stress or test in a pressure chamber. Container 

weight decreases between irrigations because the water 

in the growing medium is lost through evaporation and 

transpiration. When the container weight reaches some 

predetermined level, the crop is irrigated. With experience, workers can develop an intuitive sense of this level 

based on picking up a few randomly spaced trays (figure 

11.15C). It can also be done objectively, weighing containers on a simple household scale (figure 11.15D).

The procedure is to completely saturate the growing media in a container or block of containers and let 

it drain—measured as the “wet weight.” The container 

or block is then sown as usual, marked with a tag or flag 

stake, and placed out in the nursery. Then, once or twice 

per week during the growing season, this monitoring 

block is weighed and the weight is recorded as a percentage of the wet weight. A typical weight chart is shown in 

table 11.4 with the decision point on when to water at 85 

percent of the wet weight during the rapid growth phase 

or 70 percent during the hardening phase. Decision points 

will vary between species because of the physiological 

response of different species to moisture stress.

Assessing Irrigation Distribution

Irrigation systems need to be checked every few 

months because nozzles or drippers can become plugged 

or wear down to the point that they are no longer operating 

properly. The evenness of overhead irrigation systems can 

be easily checked by running a simple “cup test,” which 

involves measuring the irrigation water caught in a series 

of cups laid out on a regular grid system throughout the 

growing area (figure 11.16). Containers for cup tests 

should have circular openings that have narrow rims; the 

shape of the container below the opening is not important as long as the cups are stable and 2 to 4 in (5 to 10 

cm) deep to hold water without any splashing out.

Simply distribute empty cups evenly throughout the 

nursery irrigation station and run the overhead irrigation as usual for a standard time period. Turn off the system and measure the depth of the water in the cups. If 

water depth is not relatively uniform among cups, check 

pressure in the line and also check for clogs or problems 

with individual nozzles.

Types of Irrigation Systems

The best method of applying irrigation water in tropical nurseries depends on the water requirements of the 

species being grown and on the size and complexity of the 

operation (table 11.5). Small nurseries and those growing a 

variety of species may prefer hand watering for irrigation. 

Large nurseries growing only a few species may use some 

sort of mechanical irrigation system. Most nurseries use a 

combination of several systems in different watering zones 

to fulfill their irrigation needs (figure 11.17).

With the exception of hand-watering, most irrigation 

systems can be hooked up to automatic controllers based 

on timers or container weight so that irrigation can be 

automatically applied. Controllers enable the nursery manager to preprogram periods of irrigation, thereby saving 

time and labor. The prudent grower, however, will never 

220 Tropical Nursery Manual

Table11.4—An example of a weekly irrigation-monitoring program using container weights. Adapted from Dumroese and others (2008).

Visual and Tactile Clues for 

Monitoring Irrigation

• Leaves should look and feel firm, not wilted.

• Potting medium should be moist throughout the 

plug; moisture should come out when squeezed.

• Containers should feel relatively heavy when lifted.

• Immediately after watering, some water will drip 

(but not stream) out of the bottom of each container 

to indicate plants were irrigated sufficiently.

Mon Tue Wed Thu Fri Sat Sun

Wet weight 26.0 26.0 26.0 26.0 26.0 26.0 26.0

Actual weight 22.0 25.0 23.5 21.0 24.5 21.5 25.0

Wet weight (%) 85 96 90 81 94 83 96

Need to water? Yes No No Yes No Yes No
become completely reliant on automatic systems and will 

continue to directly monitor irrigation efficiency and its 

effect on plant growth on a regular basis.

Hand Watering

Hand watering is often the most practical irrigation 

strategy for small tropical plant nurseries, nurseries producing a wide diversity of species with radically different water 

requirements, or nurseries in the startup phase. Hand watering requires simple and inexpensive equipment; a hose, a 

couple of different nozzle types, and a long-handled spray 

wand are all that are absolutely necessary. The watering job 

will be more pleasant and efficient with a few additional small 

investments, such as overhead wires to guide the hoses and 

rubber boots for the staff (Biernbaum 1995). Although the 

task may appear easy, good technique and the application 

of the proper amount of water to diverse species of plants 

in different containers and at different growth stages is very 

challenging. Nursery managers need to ensure that irrigators have a conscientious attitude and are properly trained to 

work effectively with water application.

Water Quality and Irrigation 221

Figure 11.16—Periodic checks of water distribution can easily be done with a “cup test” in which cups are arranged in a grid pattern (A). 

The depth of water in the cups is measured after a standard watering period (B). Photo A by Kim M. Wilkinson, and photo B by Thomas D. 

Landis.

Figure 11.17—Most native plant nurseries use a combination of 

irrigation types to meet the needs of diverse species. This growing 

area of a nursery has both overhead sprinklers and access to a hose 

for hand watering. Photo by Thomas D. Landis.

A B
222 Tropical Nursery Manual

Advantages Disadvantages

Hand watering

• Requires inexpensive equipment that is simple to install

• Is flexible and can adjust for different species and container sizes

• Irrigators have a daily connection to the crop and can 

scout out diseases or other potential problems

• Allows water to be directed under plant foliage, reducing 

risk of diseases

• Is time consuming and labor intensive

• Involves a daily responsibility including weekends and holidays

• Requires skill, experience, and presence of mind to do properly

• Presents a risk of washing out or compacting growing medium

Microirrigation

• Water is delivered directly to the root zone of plants 

(not to foliage, where it may cause disease)

• Use of water is very efficient; less than 10 percent of applied 

water is wasted

• Delivery is uniform; an even amount of water is applied to 

each container

• Infiltration rate is good (because of slow delivery).

• The amount of leachate is low

• Designing the system and installing individual emitters for 

each plant is difficult and time consuming

• It is not generally efficient to install for plants grown 

in containers smaller than 1 gallon in size

• Each irrigation station must run a long time because of slow 

water delivery

• Emitters can plug easily (water filtration and frequent 

irrigation system maintenance is required)

• With drippers, it is difficult to verify water delivery visually; 

often, problems are not detected until it is too late

Sprinkler irrigation

• Relatively simple and inexpensive to design and install

• A variety of nozzle patterns and application rates are available

• Water distribution patterns can be measured with a cup test

• Foliar interception makes overhead watering ineffective for 

large-leaved crops

• Irrigation water can be wasted because of inefficient 

circular patterns

• An increased risk of foliar diseases is possible from excessive 

water on leaves

• For overhead sprinklers, nozzle drip from residual water in lines 

can harm germinants and young plants

• For basal sprinklers, irrigation lines must run along the floor, 

creating obstacles for workers and equipment

Subirrigation

• Although commercial products are available, subirrigation 

systems can be constructed from affordable, local materials

• Foliage remains dry, reducing the risk of foliar diseases

• Water use is efficient (up to 80 percent less water use than 

overhead watering systems)

• Application among plants is uniform

• Lower fertilizer rates are possible

• Reduced leaching of mineral nutrients is possible

• Drainage water can be captured for reuse or recycling

• No splashing disrupts or displaces mulch, germinants, 

or medium

• Provides the ability to irrigate different size containers and 

different age plants concurrently

• Is efficient in terms of time and labor requirements 

following installation

• Overhead or hand watering may be required to ensure 

sufficient surface moisture until seeds germinate

• It cannot be used with poor-quality water because salt 

buildup would occur

• Less air pruning of roots occurs

• Risk of spreading waterborne diseases is greater

• High humidity within plant canopy is possible

Table11.5—Types of irrigation systems for container nurseries.
Good hand-watering practices include the following—

• Direct water to the base of the plants.

• Avoid spraying the foliage to conserve water and 

preclude foliar diseases.

• Angle the watering nozzle down (not sideways) 

and keep pressure low to avoid washing out seeds, 

medium, or mulch.

• Use an appropriate nozzle type and water volume for 

each crop: a very fine, gentle spray for young germinants and a larger volume nozzle for larger plants.

• Adjust the flow, volume, and pace of watering to irrigate efficiently without wasting water or compacting 

medium (Biernbaum 1995).

• Achieve uniformity of water distribution so all plants 

are well irrigated; account for microclimate differences in nursery. For example, plants on the outer 

edge of a bench or in direct sun will need more water 

(figure 11.18).

• Attend to the individual needs of each crop and 

its development phase so none are overwatered or 

underwatered; develop a feel for the watering needs of 

crops over time.

Microirrigation

For nurseries that grow plants in 1-gal (4-L) or larger containers, microirrigation can be a very efficient method for 

water delivery. Microirrigation usually involves polyethylene 

pipe fitted with microsprayers (sometimes called “spitters” or 

“spray stakes”) (figure 11.19A), drippers inserted individually 

into each container (figure 11.19B), or smaller lateral tubing 

to reach all areas on the bench (figure 11.19C). Microsprayers are often preferred to drippers because they wet more 

surface area and distribute water more evenly throughout the 

container. It is also easier to visually verify the operation of a 

sprayer than a dripper. Filtration is a necessity for microirrigation systems to prevent emitters from clogging. Because 

of the slow infiltration rate of microirrigation systems, each 

irrigation station will need to run a long time to deliver adequate water to plants. Also, if containers are allowed to dry 

Water Quality and Irrigation 223

Figure 11.18—Nursery plants on the outside of a bench or 

table always dry faster, especially in windy weather. Photo by 

Thomas D. Landis.

Figure 11.19—Spray stakes are effective only for larger containers and work well because you can see them functioning and they 

have more even distribution (A). Drip emitters can also be used for larger containers (B, C). Illustration A from Dumroese and others 

(2008), photo B by Thomas D. Landis, and photo C by Brian F. Daley.

A B C
out, hand watering may be necessary to rewet the growing 

medium before drip irrigation will work.

Fixed Irrigation Systems

Overhead sprinkler systems are a common type of irrigation system and the kind that many people imagine when 

they think of irrigation. Many types of overhead irrigation 

systems exist, ranging from fixed sprinklers to moving 

boom systems.

Fixed Overhead Sprinklers

Fixed overhead sprinkler systems consist of a series 

of parallel irrigation lines, usually constructed of plastic 

polyvinyl chloride (PVC) pipe, with sprinklers spaced at 

uniform intervals to form a regular grid pattern. Overhead 

sprinklers apply water at a fairly rapid rate and will do an 

acceptable job if properly designed and maintained.

Several types of spray nozzles are used for fixed 

overhead irrigation systems. Spinner sprinklers, which 

have offset nozzles at the end of a rotating arm, spin in 

a circle when water pressure is applied (figure 11.20A). 

Stationary nozzles (figures 11.20B, 11.20C) have no moving parts but distribute water in a circular pattern; these 

nozzles also come in one-half and one-fourth of a circle 

patterns (figure 11.20D) so that full overlap coverage 

can be obtained by placing irrigation lines around the 

perimeter of the area to be irrigated. Mist nozzles are 

also sometimes installed on overhead irrigation lines and 

are primarily used during the germination period and for 

cooling and humidity control.

In general, the propagation environment is divided 

into irrigation “bays” or “zones” depending on the number of nozzles that the pump can operate at one time at 

the desired water pressure. Ideal operating pressures vary 

with the type of sprinkler, and specifications are available 

224 Tropical Nursery Manual

Figure 11.20—Either rotating spinners (A) or stationary nozzles (B) can be used with overhead (C) or basal sprinkler systems. 

In addition to the full-circle version, stationary nozzles are available in one-fourth, one-half, and three-fourths circle versions (D). 

Irrigation can be controlled with solenoid valves (E) connected to timers. Photos A and B by Thomas D. Landis, photo C by Diane 

L. Haase, illustration D by Jim Marin, and illustration E by John W. Bartok, Jr.

B

A C E

D
from the manufacturer. A solenoid valve control (figure 

11.20E) in each bay can be connected to an irrigation 

timer so that the duration and sequence of irrigation can 

be programmed. The size of each irrigation bay can be 

designed to accommodate plants with differing water 

requirements within a larger growing structure. When 

designing a new irrigation system, it is a good idea to 

obtain the help of an irrigation specialist to ensure that 

the system has balanced coverage and water pressure.

Fixed Basal Sprinklers

Basal irrigation systems are commonly used in large 

outdoor growing or holding areas (figures 11.21A, 11.21B). 

They are similar to overhead systems in design and operation in that they use a regular grid of permanent or movable irrigation lines with regularly spaced sprinklers. Both 

stationary sprinklers and rotating-impact nozzles (figures 

11.21B, 11.21C) are commonly used. These sprinklers 

rotate slowly because of the impact of a spring-loaded 

arm that moves in and out of the nozzle stream. Rotatingimpact sprinklers are available from several manufacturers in a variety of nozzle sizes and coverage capabilities. 

Because the water pressure out of the nozzle jet drives the 

impact arm, the water distribution pattern of these sprinklers is particularly dependent on proper water pressure. 

One advantage of basal irrigation systems is that impact 

sprinklers have relatively large coverage areas, which 

means fewer nozzles and less irrigation pipe are required.

Designing and Monitoring Fixed 

Sprinkler Systems

The efficiency and uniformity of an irrigation system 

is a function of five factors: (1) nozzle design, (2) size 

of the nozzle orifice, (3) water pressure and application 

rate at the nozzle, (4) spacing and pattern of the nozzles, 

and (5) amount of wind. Few operational procedures can 

improve a poorly designed system. Therefore, it is important to consult an irrigation engineer during the planning 

stages. Basic engineering considerations, such as friction 

loss in pipes or fittings and the effect of water pressure on 

sprinkler function, must be incorporated into the irrigation system design.

The size of the sprinkler nozzle and its resultant coverage 

pattern can be determined by consulting the performance 

specifications provided by the sprinkler manufacturer. 

Nursery managers need to select a nozzle size that is coarse 

enough to penetrate the plant’s foliage and minimize wind 

drift but not large enough to create splash problems. Most 

stationary sprinklers throw water in a circular distribution 

pattern (figure 11.22A), so irrigation systems need to be 

designed to provide adequate overlap between sprinklers 

(figure 11.22B). This consideration is especially important in shadehouses or outdoor growing areas where wind 

drift can be a problem. Too often, sprinklers are spaced at 

greater intervals in a cost-saving effort, but this practice is 

false economy considering the profound effect of water and 

injected nutrients on plant growth.

Water Quality and Irrigation 225

Figure 11.21—Fixed basal irrigation systems (A) are often used in outdoor growing areas. Rotating-impact sprinklers (B, C) are 

commonly used because of their greater coverage. Photo A by Brian F. Daley, and photos B and C by Thomas D. Landis.

A B C
Because water pressure has such an effect on sprinkler 

function and efficiency, it needs to be monitored regularly with a gauge permanently mounted near the nozzles 

(figure 11.22C) or with a pressure gauge equipped with a 

pitot tube directly from the sprinkler nozzle orifice. The 

pressure needs to be checked at several different nozzles 

including the nozzle farthest from the pump. The importance of regular water pressure checks cannot be overemphasized because many factors can cause a change in 

nozzle pressure. When water pressure is either too high 

or too low, it can cause erratic stripe or doughnut-shaped 

distribution patterns (figure 11.22D). Performance specifications for sprinklers at standard water pressures can 

be obtained from the manufacturer.

Moveable Boom Irrigation Systems

The most expensive but efficient type of overhead sprinkler irrigation is the moveable boom (figure 

11.23A), which applies water in a linear pattern (figure 

11.23B). Moveable booms are generally considered too 

expensive for smaller plant nurseries but need to be considered if appropriate to your scale. For more information, see Landis and others (1989).

Subirrigation

Overhead irrigation systems have often been the 

choice of container nurseries because the systems are 

relatively cheap and easy to install. The inherent inefficiency of overhead systems, however, becomes a very 

serious problem with many tropical species, especially 

those with broad leaves. Wide leaves combined with the 

close spacing of plants in a nursery create a canopy that 

intercepts most of the water applied through overhead 

irrigation systems, reducing water use efficiency and 

creating variable water distribution among plants (figures 11.24A, 11.24B). These problems can be precluded 

by subirrigation systems, which offer a promising alternative for tropical plant nurseries (Schmal and others 

2011).

Subirrigation is a relatively new irrigation option but 

has been successfully used to grow many native plants, 

from forbs (Pinto and others 2008) to trees (Dumroese 

and others 2006, Davis and others 2008). Studies taking 

place with native tropical trees and other native species 

are demonstrating that subirrigated plants can grow as 

well as plants irrigated with overhead irrigation systems, 

but with less water use, less water wasted, and less run226 Tropical Nursery Manual

Figure 11.22—Because sprinklers produce a circular irrigation pattern (A), proper spacing is critical to produce enough overlap 

(B). Check the water pressure of irrigation systems annually to ensure efficient nozzle operation (C) before irrigation problems 

become apparent (D). Photos A, C, and D by Thomas D. Landis, and illustration B by Jim Marin.

A B C 

D
off of nitrogen fertilizers (Dumroese and others 2007). 

In subirrigation systems, the bottoms of containers are 

temporarily immersed in water on a periodic basis (for 

example, for several minutes once a day). The water is 

then drained away, leaving the growing medium thoroughly wet while the leaves remain dry.

Subirrigation systems rely on capillary action to move 

water up through the growing medium against gravity. 

Capillarity is the result of the attraction of water molecules for each other and other surfaces. After the subirrigation tray is flooded, water will move up through the 

growing medium in the containers (figure 11.24C). The 

height to which water will move will depend on the characteristics of the container and the growing medium. The 

smaller the pores between the growing medium particles, 

the higher the water will move.

Several different subirrigation systems have been 

developed. Some, such as capillary beds and mats, will 

not work with the narrow-bottomed containers often 

used in tropical nurseries. With ebb-and-flow (or ebband-flood) subirrigation systems, containers sit in enclosures mounted on benches or on the floor. The enclosure 

may be a bench (figures 11.24D, 11.24E) or tray (figure 

11.24 F) designed for subirrigation or may be constructed 

from pond liner material surrounded by a raised border 

of wood or masonry. Another type of subirrigation is 

the flooded floor type, where special concrete flooring 

is flooded and then water is drained. In trough subirrigation systems, enclosures are mounted on benches at 

a slight slope, and water is flowed through the enclosures. The drained water can be captured in a holding 

tank and reused on the crops or directed to landscaping, 

stooling beds, or other parts of the nursery, or drained 

to the ground (use caution with this step to ensure fertilizer salts do not affect groundwater quality). If water is 

recycled, be sure to monitor water quality periodically to 

avoid possible problems with accumulated salts or disease transfer.

Subirrigation systems are ideal for many topical 

native plants. Although prefabricated subirrigation systems are available commercially, nurseries on a limited 

budget may consider designing their own systems using 

available materials. For example, systems can be made 

out of concrete blocks and pond liner or out of prefabricated drainable plastic ponds, even plastic “kiddie pools” 

(figure 11.24 G) (Schmal and others 2007). (Note: Some 

materials, such as galvanized metal, are inappropriate 

because of zinc toxicity.)

Some important design considerations exist for subirrigation systems. The drainage holes at the bottom of 

the containers must have good contact with the water for 

the water to enter the container. Subirrigation may be less 

effective during the establishment phase, when growing 

medium in the upper portions of containers needs to be 

moist to promote germination and early growth; therefore, supplemental hand watering or sprinkler irrigation may be necessary initially. Air root pruning may be 

reduced with subirrigation, resulting in a need for hand 

pruning; making this system inadvisable for use with 

plants that are very sensitive to root pruning.

Water Quality and Irrigation 227

Figure 11.23—Moveable boom irrigation systems (A) apply 

water in a very efficient linear pattern (B) but may be considered 

too expensive for many tropical nurseries. Photos by Thomas D. 

Landis.

B

A
228 Tropical Nursery Manual

Figure 11.24—Overhead irrigation is ineffective for broadleaved plants because so much water is intercepted by the foliage, called 

“the umbrella effect” (A), which can begin even when seedlings are young (B). Subirrigation works because water is drawn upward 

into the containers by capillarity (C). Subirrigation has been used effectively in a number of tropical nurseries and can be accomplished via benches (D, E), trays (F) or other enclosures (G). Illustrations A and C from Dumroese and others (2008), photos B, D, 

and E by Thomas D. Landis, and photos F and G by Diane L. Haase

B

E

C

F

A

D

G
Water Conservation and 

Managing Nursery Wastewater

Depending on the efficiency of the irrigation system, 

nursery runoff and wastewater may be important factors 

to consider. Overhead sprinkler irrigation is less efficient 

than a boom system. Microirrigation or subirrigation 

systems are much more efficient but are impractical for 

some types of containers and plants.

The problem of poor irrigation efficiency involves 

more than simply wasted water, although that alone is 

a concern both environmentally and economically. In 

addition, many container nurseries apply some or all of 

their fertilizer and pesticides through the irrigation system. Liquid fertilizer is usually applied in excess of the 

actual amount needed to saturate the growing medium 

and to leach excess salts. Most pesticides are applied in a 

water-based carrier. Some of these chemicals inevitably 

end up in the nursery’s wastewater runoff.

Until recently, it was thought that the soil beneath 

the nursery filtered out or decomposed fertilizer salts 

and pesticides, but this belief has been refuted. Leaching 

tests in conifer nurseries have shown that excess fertilizer nutrients and pesticides leach downward and may 

contaminate groundwater. Maximizing the efficiency of 

irrigation systems and implementing water conservation 

strategies (table 11.6, figure 11.25) is the most effective 

way to reduce runoff.

The direct recycling of used nursery water is generally not done except in subirrigation systems. Otherwise, 

on a small scale, the expense of water treatment and 

the risks of reintroducing excess salts or pests make the 

practice unpopular. Recycle systems for nursery runoff 

water may be economically viable in very water-limited 

areas, however. Less high-tech options for water reuse 

can make use of an impermeable nursery floor (such as 

pond liner) to collect water runoff from the nursery. This 

water can be stored in a pond or tank or run directly to 

other crops. Crops that are more tolerant of salts, such 

as rushes, may benefit from using runoff water, and 

these crops will even clean and filter the water. Nurseries growing aquatic or semiaquatic plants may be able 

to direct runoff to these plants and thereby increase the 

water-use efficiency of the nursery operation. Crops in 

the field, such as seed orchards, wetland crops, surrounding landscaping, or banana, palm, or tree crops, can all 

benefit from being irrigated with nursery runoff water. If 

the crops are located downhill from the nursery itself, the 

system can be gravity fed. Otherwise, a pump will be necessary to apply the water to the crops.

Water Quality and Irrigation 229

Table 11.6—Nurseries can incorporate a variety of horticultural practices to use irrigation water effectively and efficiently.

Nursery practice Conservation effect

Mulches (figure 11.25A) Reduces evaporation from the surface of the growing medium; 

the larger the container, the greater the savings

Windbreaks (chapter 2, figure 2.13A) Reduces water loss and seedling stress because of wind

Remove cull plants and minimize space between 

containers, including aisles Reduces unnecessary use of water and resultant runoff

Shadecloth and shadehouses (figure 11.25B) Reduces water use for species that do not require full sunlight

Catch runoff and recycle water for use on landscaping and 

other crops Saves considerable amounts of water and reduce fertilizer use

Treat runoff water via filtration or bioswales Reduces contaminants in recycled water
References

Biernbaum, J. 1995. How to hand water. Greenhouse Grower. 

13(14): 39, 24, 44.

Cayanan, D.F.; Zheng, Y.; Zhang, P.; Graham, T. 2008. Sensitivity 

of five container-grown nursery species to chlorine in overhead 

irrigation water. HortScience. 43: 1882–1887.

Davis, A.S.; Jacobs, D.F.; Overton, R.P.; Dumroese, R.K. 2008. 

Influence of irrigation method and container type on growth of 

Quercus rubra seedlings and media electrical conductivity. Native 

Plants Journal. 9: 4–12.

Dumroese, R.K.; Jacobs, D.F.; Davis, A.S.; Pinto, J.R.; Landis, T.D. 

2007. An introduction to subirrigation in forest and conservation 

nurseries and some preliminary results of demonstrations. In: 

Riley, L.E.; Dumroese, R.K.; Landis, T.D., eds. national proceedings: forest and conservation nursery associations—2006. 

Proceedings RMRS-P-50. Fort Collins, CO: U.S. Department of 

Agriculture, Forest Service, Rocky Mountain Research Station: 

20–26.

Dumroese, R.K.; Pinto, J.R.; Jacobs, D.F.; Davis, A.S.; Horiuchi, 

B. 2006. Subirrigation reduces water use, nitrogen loss, and moss 

growth in a container nursery. Native Plants Journal. 7(3): 253–261.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Fawell, J.; Bailey, K.; Chilton, J.; Dahi, E.; Fewtrell, L.; Magara, 

Y. 2006. Fluoride in drinking water. World Health Organization. London, United Kingdom: International Water Association 

(IWA) Publishing. 134 p.

Gingerich, S.B. 2003. Hydrologic resources of Guam. WaterResources Investigation Report 03-4126. Honolulu, HI: U.S. 

Department of the Interior, Geological Survey. 3 p.

Handreck, K.A.; Black, N.D. 1984. Growing media for ornamental plants and turf. Kensington, Australia: New South Wales University Press. 401 p.

230 Tropical Nursery Manual

Figure 11.25—Water can be conserved by many practices, such as covering sown containers with a mulch (A); reducing transpiration 

and evaporation water loss with shade (B); and insulating crop areas, such as with straw bale insulation around the perimeter of the 

growing area (C). Illustration A from Dumroese and others (2008), photo B by Thomas D. Landis, and photo C by Tara Luna.

B

C

A
Jones, J.B., Jr. 1983. A guide for the hydroponic and soilless culture grower. Portland, OR: Timber Press. 124 p.

Kaapuni Nursery. 2011. Cordyline terminalis ‘Ti’ plant. Kapahi, 

Kaua‘i, HI: Kaapuni Nursery. http://www.kaapuninursery.com/

Nursery.html. (December 2011).

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1989. 

The container tree nursery manual: volume 4, seedling nutrition 

and irrigation. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 119 p.

Pinto, J.R.; Chandler, R.; Dumroese, R.K. 2008. Growth, nitrogen 

use efficiency, and leachate comparison of subirrigated and overhead irrigated pale purple coneflower seedlings. HortScience. 43: 

897–901.

Robbins, J. 2011. Irrigation water for greenhouses and nurseries. 

Publication FSA6061. Little Rock, AR: University of Arkansas, 

Horticulture Department. 6 p.

Schmal, J.L.; Wollery, P.O.; Sloan, J.P.; Clark, D.F. 2007. A 

low-tech, inexpensive subirrigation system for production of 

broadleaved species in large containers. Native Plants Journal. 

8: 267–269.

Schmal, J.L.; Dumroese, R.K.; Davis, A.S.; Pinto, J.R.; Jacobs, D.F. 

2011. Subirrigation for production of native plants in nurseries – 

concepts, current knowledge, and implementation. Native Plants 

Journal. 12:81-93.

Stetson, L.E.; Mecham, B.Q. 2011. Irrigation. 6th ed. Falls Church, 

VA: Irrigation Association. 1,089 p.

Tchobanoglous, G.; Schroeder, E.D. 1985. Water quality: characteristics, modeling, modification. Melo Park, CA: Addison-Wesley 

Publishing Co. 768 p.

Zheng, Y.; Cayanan, D.F.; Dixon, M. 2009. Control of pathogens 

in irrigation water using chlorine without injury to plants. International Plant Propagators’ Society, Combined Proceedings. 58: 

248–249.

Water Quality and Irrigation 231
232
233

Plants require adequate quantities of mineral nutrients in the proper balance for 

basic physiological processes, such as photosynthesis, and to promote rapid growth 

and development. Without a good supply of mineral nutrients, growth is slowed and 

plant vigor reduced. Young plants rapidly deplete mineral nutrients stored within 

their seeds, and cuttings have limited nutrient reserves. Therefore, nursery plants 

must rely on root uptake of nutrients from the growing medium. When nutrients 

are supplied in proper amounts and at the proper time, nursery plants can achieve 

optimum growth rates.

This chapter describes the importance of nutrition to plant growth development 

and details typical fertilization practices for producing plants in tropical nurseries.

Plant Nutrition and Fertilization

Douglass F. Jacobs and Thomas D. Landis 12

Facing Page: Topdressing native sandalwood (Santalum species) seedlings with controlled-release fertilizer at Waimea State 

Tree Nursery in Hawai‘i. Photo by Douglass F. Jacobs.
Basic Principles of Plant Nutrition

A common misconception is that fertilizer is “plant food” 

(figure 12.1A), but the basic nutrition of plants is very different from that of animals. Using the green chlorophyll in their 

leaves, plants make their own food in the form of carbohydrates from sunlight, water, and carbon dioxide in a process 

called photosynthesis (figure 12.1B). These carbohydrates 

provide the plant energy, and when combined with mineral 

nutrients absorbed from the soil or growing medium, carbohydrates are used to synthesize proteins and other compounds necessary for basic metabolism and growth.

Thirteen mineral nutrients are considered essential to 

plant growth and development and are divided into macronutrients and micronutrients based on the amounts found 

in plant tissue (table 12.1). Some mineral nutrients have a 

structural function. For example, nitrogen is an integral 

part of plant proteins, and nitrogen and magnesium are 

structural components of chlorophyll molecules needed 

for photosynthesis (figure 12.2). Understanding these basic 

functions has practical applications because a deficiency of 

either nutrient causes plants to be chlorotic (that is, yellowish in color). Other mineral nutrients have no structural 

role, but potassium, for example, is functionally important 

in causing the stomata in leaves to open and close.

234 Tropical Nursery Manual

Figure 12.1—Although some fertilizers are advertised as “plant food” (A), plants actually create their own food through the process 

of photosynthesis in their green leaves (B). Photo A by Thomas D. Landis, and illustration B by from Dumroese and others (2008).

Figure 12.2—Mineral nutrients such as nitrogen and magnesium 

are important components of chlorophyll molecules that give 

leaves their green color and are essential for photosynthesis. 

Illustration from Dumroese and others (2008).

A B
Name

 (symbol)

Percent of 

plant tissue

(oven dry 

weight)

Structural 

functions

Physiological

functions

Deficiency symptoms

(general)

Macronutrients

Nitrogen

(N)

1.5 Component of 

chlorophyll, amino acids, 

proteins, and nucleic acids

Vital to all physiological processes 

through roles in enzyme systems.

Old leaves turn yellow (chlorosis), plant 

growth stunted; small leaves. Differs 

from iron chlorosis because older foliage 

is affected first.

Phosphorus

(P)

0.2 Component of cell walls 

and nucleic acids

Energy storage and release, 

functioning of cell membranes, 

regulates enzyme and cell buffering.

Some plants show purpling, but stunting 

is most common.

Potassium

(K)

1.0 None Osmotic adjustment. Important in 

maintaining cell turgor, phloem 

transport, cell growth.

Necrosis of leaf margins, stunting.

Calcium

(Ca)

0.5 Component of 

cell walls

Facilitates cell division, stabilizes cell 

membranes, inhibits fungal infections 

and toxins.

Disintegration of cell walls, inhibition 

of meristems and especially root tips, 

increased susceptibility to fungal attack.

Magnesium 

(Mg)

0.2 Component of 

chlorophyll

Regulates cellular pH and ionic balance, 

energy transfer, enzyme stabilization.

Interveinal chlorosis or yellowing of 

needle tips, symptoms may occur in 

patches.

Sulfur 

(S)

0.1 Maintain protein 

structure, component 

of vitamins, and 

coenzyme A

Amino acids formation, protein 

synthesis, ion transport across 

membranes.

Light green foliage leading to chlorosis, 

often visible on younger foliage, stunting.

Micronutrients

Iron

(Fe)

0.01 Critical to manufacture 

of chlorophyll 

Catalyst to several enzymes systems, 

especially for photosynthesis.

Chlorosis of newer foliage; in severe 

cases leaves turn whitish.

Manganese 

(Mn)

0.005 Structural component 

of ribosomes

Critical to Hill Reaction in photosynthesis, carbohydrate and nucleic acid 

synthesis, and lipid metabolism.

Chlorosis similar to iron deficiency 

except severe cases lead to necrosis.

Zinc

(Zn)

0.002 Component of several 

enzymes

Enzyme catalyst for carbohydrate 

metabolism, protein synthesis, and 

auxin production.

Reduced internode elongation (“rosetting”) and stunted foliage (“littleleaf ”). 

Chlorosis or bronzing of younger leaves.

Copper

(Cu)

0.0006 Constituent of proteins 

and enzymes

Photosynthetic functioning; phenol, 

carbohydrate, and nitrogen metabolism.

Chlorosis, tip dieback, twisted, 

needle tips.

Molybdenum 

(Mo)

0.00001 Component of several 

enzyme systems, notably 

for nitrogen uptake

Essential for reducing nitrate ions to 

ammonium after uptake; critical for 

nodule function in legumes.

Chlorosis and necrosis of leaf tissue.

Boron 

(B)

0.002 None Involved in cell division and 

elongation, lignification of cell walls, 

synthesis of amino acids and proteins.

Distortion and discoloration of shoot and 

root tips, short shoot internodes producing a bushy or rosette appearance.

Chloride 

(Cl)

0.01 None Osmosis and ion balance within cells; 

photosynthetic oxygen evolution.

Rare. Wilting of leaf margins and 

restriction of root elongation.

Table 12.1—The 13 essential plant nutrients (divided into macronutrients and micronutrients). Nitrogen, phosphorus, and potassium 

are the most common fertilizer elements.

Plant Nutrition and Fertilization 235
An important nutrition concept is Liebig’s Law of the 

Minimum, which states that plant growth is controlled 

by the mineral nutrient in shortest supply. See Chapter 5, 

Propagation Environments, for more discussion about limiting factors. A good way to visualize the concept of limiting 

factors is a wooden bucket with staves of different lengths. 

If water is poured into the bucket, it can be filled only to the 

height of the shortest stave—the limiting factor. Nitrogen 

is nearly always limiting to plant growth in nature, which 

is the reason why nitrogen fertilizer is frequently applied in 

nurseries (figure 12.3).

Just as important as the absolute quantities of nutrients 

available to plants is the balance of nutrients. The proper 

nutrient balance is relatively consistent among plant species and can be expressed as Ingestad’s Ratios. Healthy 

plant tissue contains approximately 100 parts of nitrogen 

to 50 parts of phosphorus, to 15 parts of potassium, to 5 

parts of magnesium, to 5 parts of sulfur. On a practical 

basis, most tropical plant nurseries use complete fertilizers 

that contain a balance of all mineral nutrients.

Mineral nutrients are absorbed by root hairs as two 

types of ions: cations and anions. Cations have an electrically positive charge, while anions are negatively charged. 

Particles of soil, compost, and artificial growing medium 

are also charged, so nutrient ions attach to sites with an 

opposite charge. Roots typically release a cation (often 

H+) or an an ion (for instance, HCO3

–) when taking up a 

nutrient ion of the same charge from the soil or growing 

medium (figure 12.4). See Chapter 6, Growing Media, for 

more information on this topic. Fertilizers applied in the 

nursery break down into soluble nutrient ions, which are 

then taken up by plant roots.

Sources of Mineral Nutrients

Plants produced in container nurseries may acquire 

nutrients from several different sources, including the 

growing medium, irrigation water, beneficial microorganisms, and fertilizers. Many tropical nurseries use artificial 

growing media that are essentially infertile, which enables 

growers to apply the right type of fertilizer, in the right 

amount, and at the right time. Levels of mineral nutrients in a commercial growing medium are generally very 

low, which is why they are often amended with a “starter 

dose” of fertilizer. Native soils and composts contain higher 

nutrient concentrations than commercial growing media 

but rarely enough for the fast growth rate and nutrient balance desired in nurseries. If soil or other homemade growing media will be used, a soil test can be done to determine 

which nutrients are lacking (see the section on testing in 

the following paragraphs).

236 Tropical Nursery Manual

Figure 12.3—The concept of limiting factors can be illustrated by a wooden bucket that can be filled with water only to the 

shortest stave. In this example, nitrogen (N) is the most limiting, 

which is typical in nurseries. Illustration from Dumroese and 

others (2008). 

Figure 12.4—Nutrient cations and anions are extracted by plant 

roots through an exchange process with soil, compost, or artificial 

growing media. Illustration from Dumroese and others (2008).
Another potential source of mineral nutrients in nurseries is irrigation water. “Hard water” often contains 

enough calcium and magnesium for good plant growth. 

Sulfur is another nutrient often found in water sources. 

Beneficial microorganisms may provide an important 

source of nitrogen for some species, such as legumes, as 

detailed in Chapter 13, Beneficial Microorganisms.

To achieve the desired plant growth and health, fertilizers are the most common source of mineral nutrients 

in tropical plant nurseries. Growers need to have a good 

understanding of types of fertilizers and how and when to 

apply them. Many different types of fertilizers are used in 

tropical plant nurseries and vary according to their source 

materials, nutrient quantities, and mechanisms of nutrient 

delivery. The two main fertilizer categories are organic (figure 12.5A) and synthetic (figure 12.5B).

All commercial fertilizers are required by law to show 

the ratio of nitrogen to phosphorus to potassium (actually 

the oxides of phosphorus and potassium; N:P2O5:K2O) and 

the complete nutrient analysis on the label (figure 12.5C). 

Some fertilizers contain only one mineral nutrient whereas 

others contain several. Examples of single-nutrient fertilizers include ammonium nitrate (34-0-0) or concentrated 

superphosphate (0-45-0). Fertilizers may be blended or 

reacted to supply two or more essential nutrients. An example of a blended fertilizer is a 12-10-8, which was formed 

by adding triple superphosphate (0-46-0), potassium magnesium sulfate (0-0-22), and ammonium nitrate (34-0-0). 

An example of a multiple-nutrient fertilizer is potassium 

nitrate (13-0-44).

Some tropical plant nurseries prefer organic fertilizers because they are less likely to burn crops if too much 

is applied and have lower risk of water pollution. If inoculating with beneficial microorganisms, organic fertilizers 

are generally preferred. The main drawbacks of organic 

fertilizers are that they are more expensive, and their 

lower nutrient content and solubility result in slower plant 

growth. Most of the research in tropical plant nurseries has 

Plant Nutrition and Fertilization 237

Figure 12.5—The two main types of commercial fertilizers are organic (A), and synthetic (B). All fertilizers must show the 

nitrogen-phosphorus-potassium ratio on the label, and most also give a complete listing of mineral nutrients (C). Photos by Thomas 

D. Landis.

B

C

A
been done with synthetic fertilizers because they are inexpensive, readily soluble, and quickly taken up by crops. 

Synthetic fertilizers are popular with growers because they 

stimulate the rapid growth rates desired in nursery culture.

Organic Fertilizers

Until the early 20th century, nearly all fertilizers used in 

agriculture, including nurseries, were organic. The shortage 

of Chilean nitrate, a major source of organic nitrogen, led to 

development of the Haber-Bosch Process, which converts 

the abundant nitrogen gas in our atmosphere into ammonia. 

This ammonia can then be chemically converted into a vast 

array of synthetic fertilizers. After the Second World War, 

these synthetic, ammonia-based fertilizers became cheap 

and readily available, and use of organic fertilizers dropped 

from 91 percent in the early 1900s to 3 percent by the 1950s. 

In recent decades, however, organic farming has seen a resurgence because of changes in public values. In tropical areas, 

concern about the environmental and economic implications of manufacturing, transporting, and using synthetic 

fertilizers is rising. In response to this increasing demand, a 

wide variety of organic fertilizers have been developed, and 

many have potential in tropical plant nurseries.

Defining organic fertilizers can be a complicated and confusing subject, and much of this confusion comes from terminology. For our purposes, organic fertilizers can be defined 

as materials that are naturally occurring and have not been 

synthesized, and we recognize two general categories: animal 

or plant wastes and natural minerals (figure 12.6).

Animal or Plant Wastes

These materials are what most people consider to be 

organic fertilizers and can be applied to crops directly or 

developed into a wide variety of other processed fertilizers. 

One of the attractions of these types of organic fertilizers is 

they are renewable and widely available.

Unprocessed Organics

This category is by far the largest and most complicated because nearly any type of animal or plant waste can 

be used as a fertilizer, including animal manure, sewage 

sludge, and peat moss. In tropical areas, guano is an excellent organic fertilizer and consists of the accumulated 

excrement of seabirds or bats. Guano has high levels of 

phosphorus and nitrogen and does not have any noticeable odor. One of the largest mining operations occurred 

on the small South Pacific island of Nauru where centuries of deposition by seabirds created vast reserves of 

mineralized guano.

The best criterion for determining which types of 

unprocessed organic matter are suitable fertilizers is the 

carbon-to-nitrogen (C:N) ratio. Organic materials with a 

C:N less than 10:1 are considered to be fertilizers. Unprocessed organics can be challenging to use because of their 

high potential for water pollution. 

Processed Organics

This category includes any organic material that has been 

processed in some manner before being used as a solid or liquid fertilizer, and includes composts, blood meal, bone meal, 

and materials such as feather meal and kelp extracts (table 

12.2). Manure is a good source of nutrients but will always 

need to be composted first so it will not damage plants or 

pollute water. One way to test if the manure is composted 

enough to use as fertilizer is to place a few handfuls into a 

plastic bag and seal it for 24 hours; if the material does not 

smell bad or give off heat at the end of the test period, it is 

ready to use; otherwise, it will need further composting. 

Nearly any organic waste matter can be composted and the 

composting process has been well documented. See Chapter 

6, Growing Media, for more information on composting. 

Many new processed organic fertilizers are now available 

from horticultural supply sources.

238 Tropical Nursery Manual

Figure 12.6—The terminology of organic fertilizers is complicated, but various types can be illustrated in this flow chart 

from Landis and Dumroese (2011).
Anyone considering the use of processed organic fertilizers needs to test them first to decide if a need exists 

for composting, determine proper application rates, and 

identify potential nutrient toxicities or deficiencies. It is 

also wise to initiate small-scale tests with any new fertilizer product to avoid possible problems and determine 

plant growth rates.

Natural Mineral Fertilizers

This second major category of organic fertilizers includes 

minerals and other materials that come directly from the 

earth (figure 12.6). Minerals like sodium nitrate are commonly used in many blended organic fertilizers because they 

are soluble and have a high nutrient content. Like all types of 

mining, however, obtaining natural minerals is an extractive 

process and nonrenewable in the long term.

Rock Phosphate

Natural deposits of fluoroapatite are the raw material of most phosphate fertilizers and are currently mined 

in North Africa, the former Soviet Union, and in Florida, 

Idaho, Montana, Utah, and Tennessee. The raw ore contains 14- to 35-percent phosphate (P2O5) and is processed 

by grinding and washing into a fine granular fertilizer. 

Rock phosphate is very insoluble in water and makes an 

effective slow-release granular fertilizer. Because of its 

low solubility, rock phosphate has been recommended as 

an ideal phosphorus fertilizer to encourage mycorrhizal 

development.

Sodium Nitrate

This salt (NaNO3) is commonly known as Chilean or 

Peruvian saltpeter because of the large caliche mineral 

deposits found in both countries. Although this fertilizer 

has been used in organic farming for many years, several 

organic certifying agencies have concluded that mined 

mineral fertilizers conflict with basic organic principles. 

For example, the USDA National Organic Program currently restricts use of sodium nitrate to less than 20 percent 

of total annual applied nitrogen and requires that growers 

phase out its use over time.

Magnesium Sulfate

This mineral (MgSO4) comprises Epsom salts or Kieserite. Although more widely used for medicinal purposes, 

magnesium sulfate is a very soluble source of magnesium 

and sulfur and has been used in the formulation of liquid 

fertilizers for container tree nursery crops.

Sul-Po-Mag

Technically known as sulfate of potash-magnesia or 

langbeinite, this salt is mined from marine evaporite deposits and is a common component in many blended organic 

fertilizers. It was originally discovered in Germany and 

contains soluble nutrients in the following ratio: 22-percent potassium, 22-percent sulfur, and 11-percent magnesium. Another common trade name is K-Mag Natural, an 

ideal product for supplying potassium and sulfur without 

any accompanying nitrogen.

Blended Organic Fertilizers

This category of organic fertilizers includes a wide variety of products containing a mixture of processed organic 

plant or animal wastes supplemented with natural minerals (figure 12.7). It is easy to identify blended organic fertilizers by checking the ingredients on their labels.

Plant Nutrition and Fertilization 239

Organic fertilizers N P K

Bat guano (fresh) 10 3 1

Bat guano (old) 2 8 0

Blood meal 10 0 0

Bone meal (steamed) 1 11 0

Cottonseed meal 6 2 1

Eggshells 1.2 0.4 0.1

Fish emulsion 4 1 1

Fish meal 5 3 3

Greensand 0 0 7

Hoof and horn meal 12 2 0

Kelp meal 1.5 0.5 2.5

Soybean meal 7.0 0.5 2.3

Worm castings 0.5 0.5 0.3

Manure

Cow 2 2.3 2.4

Horse 1.7 0.7 1.8

Pig 2 1.8 1.8

Sheep 4 1.4 3.5

Poultry 4 4 2

Table 12.2—Percentages of nitrogen, phosphorus, and potassium 

supplied by a variety of organic materials. Adapted from Diver and 

others (2008).

N = Nitrogen, P = Phosphorus, K = Potassium
Solid Organic Fertilizers

 Powdered or granular fertilizers can be derived from 

unprocessed organics, processed organics, natural minerals, or blended organics (figure 12.6). Solid organic 

fertilizers have not been widely used in forest or native 

plant nurseries. Biosol® (6-1-3) is a solid organic fertilizer that is supplemented with Sul-Po-Mag, however, and 

has potential for nursery use.

Liquid Organic Fertilizers

This category of organic fertilizers can be derived 

from processed organics, natural minerals, or blended 

organics (figures 12.5A, 12.6). Ingredients might include 

fertilizers such as fish waste, soybean meal, kelp, recycled 

foodstuffs, bat guano, sulfate of potash, feather meal, 

blood meal, steamed bone meal, or any number of ingredients. Many products are targeted to specific crops but 

others are for more general use. Some liquid organic fertilizers contain suspended material and must be filtered 

or continually agitated during fertigation to prevent the 

material from plugging nozzles.

Synthetic Fertilizers

The most important synthetic fertilizers are made 

using the Haber-Bosch process in which atmospheric 

nitrogen is converted into ammonia under high temperature and pressure. Having been called the most 

important invention of the 20th century, this industrial 

process produces 500 million tons of artificial fertilizer 

per year. Synthetic fertilizers are popular because they 

are relatively inexpensive, readily available, and have 

high nutrient content compared with organic products. 

In populated tropical areas, synthetic fertilizers can be 

found at garden supply shops and through horticultural 

dealers, but inaccessibility and transport costs may be a 

limitation in remote areas. In the humid tropics, storage 

of synthetic fertilizers becomes a challenge because they 

readily absorb moisture from the air.

Synthetic fertilizers can be divided into two classes: 

(1) soluble products that release nutrients quickly when 

dissolved in water and (2) slow-release or controlledrelease fertilizers that release nutrients slowly over time. 

Both types have their advantages and disadvantages, 

which need to be considered before deciding upon a fertilization system (table 12.3). Other types of granular fertilizers that are used on lawns or in agriculture are not 

recommended for tropical plant nurseries.

240 Tropical Nursery Manual

Figure 12.7—Blended organic fertilizers (A) contain processed organic materials and natural minerals, such as sulfate 

of potash (B). Photos by Thomas D. Landis.

B

A
Soluble Fertilizers

Soluble synthetic fertilizers come in either granules or 

water-soluble crystals (figure 12.8A). Soluble fertilizers 

are typically injected into the irrigation system, a process 

known as fertigation (see the discussion in the following 

section). Their popularity stems from the fact that the 

application rates can be easily calculated, distribution 

is as uniform as the irrigation system, and, if properly 

formulated and applied, the chance of fertilizer burn is 

very low (table 12.3). Because they are immediately soluble, the nutrients in these fertilizers are quickly available for 

plant uptake and it is easy to adjust nutrient levels and ratios, 

which gives the grower great control of plant growth rates. 

When growing a variety of plant species in the same area, it is 

necessary to separate them based on relative growth rates so 

that different fertilizer mixes can be applied. The major drawback to using soluble fertilizers is their relatively low nutrient 

uptake efficiency and subsequent high leaching rate. Because 

they are soluble, some nutrients that are not used by plants or 

are held on the cation exchange sites of the growing medium, 

leach out with each irrigation. Fixed overhead irrigation systems apply fertilizer solution to the aisles as well as to the 

plants, so all these nutrients run off and can cause pollution.

Plant Nutrition and Fertilization 241

Factor Soluble fertilizer Controlled-release fertilizer

Nutrient release rate Very fast Much slower—dependent on type and thickness 

of coating, as well as temperature and moisture 

Number of applications Multiple—must be applied at regular 

intervals

Usually once per season, but additional top-dressing 

is an option

Uniformity of application Good, but dependent on 

irrigation coverage

Can be variable if incorporated, resulting in 

uneven growth

Adjusting nutrient rates and ratios Easy and quick Difficult

Nutrient uptake efficiency Poorer Better

Leaching and pollution potential Higher Lower

Potential for fertilizer burn 

(salt toxicity) Low if applied properly Low, unless prills damaged during incorporation 

or following high temperatures

Product cost Lower Higher

Application costs Higher Lower

Table 12.3—Comparison of advantages and disadvantages of two major types of synthetic fertilizers used in tropical plant nurseries.

A B

Figure 12.8—The two major types of synthetic fertilizers used in native plant nurseries are soluble crystals that dissolve completely in water (A), and polymer-coated controlled-release fertilizers, which are soluble fertilizers inside a thin plastic shell. These 

“prills” come in a variety of sizes, nutrient formulations, and release rates (B). Photo A by Thomas D. Landis, and photo B by 

Douglass F. Jacobs.
Special types of water-soluble fertilizers are sometimes 

applied to stimulate uptake through the foliage (“foliar 

feeding”). Although this method seems to be a good way 

to fertilize, remember that all leaves are covered with a 

water-repelling cuticle, so foliar uptake is very inefficient. 

More nutrient uptake actually occurs through the roots as 

a result of the fertilizer solution washing down into the 

soil or growing medium rather than through the foliage 

itself. Special care must be taken to prevent salt damage 

to foliage, so we do not recommend foliar feeding for 

smaller nurseries.

Controlled-Release Fertilizers

Coated fertilizers consist of a water-soluble fertilizer 

core covered with a less-soluble barrier, which affects the 

nutrient release rate. The most common coatings for controlled-release fertilizers (CRF) are sulfur or a polymer 

material. The major advantage of CRFs is that the gradual 

nutrient release results in higher nutrient uptake efficiency and less pollution potential (table 12.3). Often, CRF 

needs to be applied only once. Using CRF enables growers to supply nutrients for an extended duration without 

the specialized equipment needed to apply water-soluble 

fertilizers. Popular brands of polymer-coated fertilizers 

include Osmocote®, Nutricote®, and Polyon®, which have 

a variety of formulations for different plant species and 

growing cycles. Some release rates are so slow that they 

can provide an added benefit after outplanting.

Polymer-coated CRFs are widely used in tropical plant 

nurseries. The round, polymer-coated “prills” (figure 

12.8B) have a more uniform nutrient release than sulfurcoated products. In addition, the prills can be formulated 

to contain both macronutrients and micronutrients (table 

12.4), whereas sulfur-coated products generally supply 

only nitrogen.

Nutrient release from polymer-coated CRF is a multistep 

process. During the first irrigation, water vapor is absorbed 

through microscopic pores in the coating. This process creates an osmotic pressure gradient within the prill, causing 

the flexible polymer coating to expand. This expansion 

enlarges the tiny pores and allows the mineral nutrients to 

be gradually released into the soil or growing medium (figure 12.9). Besides water, temperature is the primary factor 

affecting the speed of this process. Nutrient release generally 

increases with increasing temperature. In warm, tropical climates, growers can expect CRFs to release more quickly than 

indicated on the label. The manufacturer adjusts the release 

rates of polymer-coated products by altering the thickness 

and nature of the polymer material, and longevities vary 

from about 3 to 18 months.

242 Tropical Nursery Manual

Nutrient Percentage

Macronutrients

 Nitrogen (7% ammonium; 8% nitrate) 15

 Phosphorus (P2O5) 9

 Potassium (K2O) 12

 Calcium 0

 Magnesium 1.3

 Sulfur 5.6

Micronutrients

 Iron (0.09% soluble, 0.01% chelated) 0.46

 Manganese 0.06

 Zinc 0.05

 Copper 0.05

 Boron 0.02

 Molybdenum 0.02

Table 12.4—Nutrient analysis of 15-9-12 Osmocote® Plus 

controlled-release fertilizer. Adapted from Everris Company 

(2012).

Figure 12.9—Nutrient release from polymer-coated fertilizers occurs after water vapor is absorbed through the prill membrane, creating an osmotic pressure gradient that expands the 

pores within the coating and allows fertilizer nutrients to pass 

through to the growing medium. Illustration from Dumroese 

and others (2008).
Other types of CRFs are nitrogen-reaction products, 

such as ureaform and IBDU Micro Grade Fertilizer. These 

fertilizers are created through a chemical reaction of watersoluble nitrogen compounds, which results in a more complex molecular structure with very limited water solubility. The rate of nutrient release of ureaform is controlled 

by many factors, including soil temperature, moisture, 

pH, and aeration, while IBDU becomes available primarily 

through hydrolysis. These materials are rarely used in tropical native plant container production but are sometimes 

applied at outplanting.

Comparison of Organic Versus 

Synthetic Fertilizers

Because of the variability involved, it is difficult to compare organic and synthetic fertilizers but some generalizations can be made (table 12.5).

Mineral Nutrient Analysis

Nearly all organic fertilizers have relatively low mineral 

nutrient analyses. The nitrogen percentage is rarely above 

15 percent and more typically in the range of 5 to 10 percent (figure 12.7B). Higher analysis products are usually 

supplemented with natural minerals such as sodium nitrate. 

Organic fertilizers often contain all 13 mineral nutrients. 

Synthetic fertilizers can be specially formulated to contain 

only a few macronutrients or a range of all mineral nutrients.

Nutrient Release Rate

One of the major differences between organic and synthetic fertilizers is how fast their nutrients become available to plants. Soluble synthetic fertilizers are formulated 

as salts and are readily available, whereas a range of release 

rates exist for synthetic CRF, depending on coating 

thickness. Unprocessed organic fertilizers must first be 

broken down into smaller particles by soil microorganisms and then converted to a soluble form. Even processed organics contain a large percentage of insoluble 

nitrogen that must undergo microbial decomposition 

before being available for plant uptake. Liquid organic 

fertilizers have the benefit of being already in solution, 

or at least, in aqueous suspension.

Compatibility With Beneficial 

Microorganisms

Perhaps one of the most underappreciated benefits of 

organic fertilizers is that they promote the growth of beneficial soil microorganisms, including mycorrhizal fungi 

and nitrogen-fixing bacteria because nutrients are released 

slowly and the organic component improves soil conditions. Research has shown that high levels of soluble synthetic fertilizers, especially nitrogen and phosphorus, can 

inhibit the establishment and development of mycorrhizal 

fungi. This is particularly evident in the soilless growing 

media of container seedlings where applications of soluble, 

synthetic fertilizers are common.

Cost

A comparison of fertilizer costs is difficult because each 

fertilizer contains different percentages of nutrients and 

values must be compared on a per-weight or per-volume 

basis. Although they can be more expensive strictly on a per 

nutrient basis, both processed and unprocessed commercial 

organic fertilizers provide many other benefits that are hard 

to valuate, including adding organic matter and stimulating 

soil microorganisms (table 12.5). Synthetic fertilizers also 

have hidden costs, such as the carbon emissions during their 

manufacture and the ecological effects of increased potential for water pollution. For tropical growers, the potential 

to reduce transport costs by sourcing local commercial or 

homemade organic fertilizers may be attractive. In the final 

analysis, fertilizers represent only a very small percentage of 

the cost of producing nursery stock, so price may not be a 

deciding factor on the type of fertilizer to use.

Handling and Application

Because of their bulkiness and low nutrient analysis, 

unprocessed organic fertilizers are more expensive to ship, 

Plant Nutrition and Fertilization 243

Factor Organic Synthetic

Mineral nutrient analysis Low High

Range of mineral 

nutrients All One to many

Nutrient release rate Slower Faster

Compatibility with 

beneficial microorganisms Yes At low levels

Cost More Less

Handling Bulkier More 

concentrated

Ecological sustainability Yes No

Water pollution risk Low High

Other benefits

Improves soil 

texture and 

encourages soil 

microbes

Better for 

research

Table 12.5—Comparison of organic and synthetic fertilizers.
store, and apply compared with synthetic fertilizers. Conversely, synthetic fertilizers are more uniform in quality, 

have a high nutrient analysis per unit weight, and are much 

easier to apply to crops.

Ecological Sustainability and Water 

Pollution

One of the real benefits of organic fertilizers is that they 

are kinder to the environment and many can be obtained 

from recycled materials. Not only can nurseries recycle 

their cull seedlings, weeds, and other organic materials 

through composting, but they can also serve to recycle 

leaves, yard clippings, and other such organic wastes from 

the local community that would otherwise go to landfills 

(Morgenson 1994).

Nutrients in organic fertilizers are much less susceptible to leaching than those in synthetic fertilizers. Both 

processed and unprocessed organic fertilizers release their 

nutrients slowly and in a form that remains in the soil profile. Synthetic fertilizers, especially soluble formulations, 

often release their nutrients much faster than plants can 

use them, with the excess nutrients potentially polluting 

surface or ground water.

Fertilizer Application Rates and 

Methods

Fertilizer application rates depend on the growing environment and other cultural factors such as container volume, 

type of growing media, and irrigation frequency. In particular, the size of the growth container has a profound effect on 

the best application rate and timing. Very small containers 

require lower rates applied frequently whereas larger containers can tolerate high application rates applied less frequentlyFor most fertilizer products, manufacturers provided 

general recommended application rates for container nursery plants on package labels. In addition, experimentation, 

consultation with other growers and development of propagation protocols (see Chapter 4, Crop Planning: Propagation 

Protocols, Schedules, and Records) will help refine fertilizer 

application rates for tropical native plant species.

Fertigation for Applying 

Soluble Fertilizers

Some tropical plant nurseries apply soluble fertilizers 

through their irrigation systems, a process known as 

fertigation. The best fertigation method varies depending 

on the type of irrigation and the size and sophistication of 

the nursery. The simplest method is to mix soluble fertilizers 

and water in a watering container or use a hose injector, and 

water plants by hand. This method can be tedious and time 

consuming, however, when fertigating a large quantity of 

plants. On the other hand, this method may be appropriate 

for smaller nurseries growing many different species with 

different fertilizer needs.

Fertilizer injectors are a much more precise way to apply 

soluble fertilizers, especially when growing large numbers 

of plants with the same fertilizer requirements. The simplest injectors are called siphon mixers (figure 12.10A) and 

the HozonTM and EZ-FLO® are common brands and range in 

cost from $20 to $200. Siphon injectors are attached to the 

water faucet and have a piece of rubber tubing inserted into 

a concentrated fertilizer solution (figure 12.10B). When an 

irrigation hose is attached to the other end and the water 

is turned on, the flow through the hose causes suction that 

pulls the fertilizer solution up and mixes it with the water 

at a fixed ratio. For example, the HozonTM injects 1 part of 

soluble fertilizer to 16 parts of water, which is a 1:16 injection ratio. Note that this injector requires a water pressure 

of at least 30 pounds per square inch (lb/in2 [psi]) (0.207 

MPa) to work properly whereas the EZ-FLO® functions at 

water pressures as low as 5 psi (0.034 MPa).

More complicated, but more accurate, fertilizer injectors 

cost from around $300 to more than $3,000. For example, 

the Dosatron® is a water pump type of injector that installs 

directly into the irrigation line and pumps the fertilizer solution into the irrigation pipe at a range of injection ratios 

(figure 12.10C). Among the most technologically advanced 

fertigation systems is the automated hydraulic boom, 

which provides very consistent and uniform coverage of 

water and fertilizer to the crop. The relatively high price 

of irrigation booms, however, makes them cost prohibitive for most tropical plant nurseries. Any injector must be 

calibrated after it is installed to verify the fertilizer injection 

ratio and then must be checked monthly to ensure that it is 

still working properly.

Some nutrients, notably calcium and magnesium, 

are very insoluble in water and can even cause solubility problems in concentrated fertilizer solutions. If they 

are not present naturally in the irrigation water, calcium 

and magnesium must be supplied in a separate fertilizer 

solution or in a dolomitic limestone amendment to the 

growing medium.

244 Tropical Nursery Manual

Caution: Every fertilizer injector must 

be installed with a backflow preventer to 

eliminate the possibility that soluble fertilizer 

could be sucked back into the water line and 

contaminate drinking water.
Applying Controlled-Release Fertilizers

CRF can be topdressed (sprinkled into the top of the 

container), if care is taken to ensure that each container 

or cell receives an equal number of prills (figure 12.11A, 

table 12.6). A special drop-type application wand can be 

used to topdress larger (>1 gal [4 L]) containers because 

a measured dose of fertilizer can be applied to the base of 

each plant. This method avoids the potential of fertilizer 

granules being lodged in foliage and burning it as soon as 

the crop is watered.

Another option for applying CRF is to incorporate it into 

the growing medium (figure 12.11B, table 12.6). If growers 

mix the growing medium on site, CRF can be incorporated but special care must be taken to ensure uniform 

distribution (figure 12.11C) and to prevent damage to 

the prill coating. If the coating is fractured, then the soluble fertilizer releases immediately, causing severe salt 

injury. Nurseries can purchase growing media with CRF 

that have already been evenly incorporated with special, commercial mixing equipment. Purchased medium 

Plant Nutrition and Fertilization 245

Figure 12.10—Soluble fertilizers can be mixed with water and 

applied to the crop, a process called “fertigation.” A siphon injector 

(A) sucks up concentrated fertilizer solution, mixes it with irrigation water, and the fertigation solution is then applied with a hose 

(B) or other irrigation system. The Dosatron® injector (C) allows 

for more precise control of injection ratios. Illustrations A and B 

courtesy of HummertTM International, and photo C by Tara Luna.

B

C

A

Units

Incorporation rates

Low Medium High

lbs per yd3 3.0 8.0 12.0

kg per m3 1.8 4.7 7.1

g per L 1.8 4.7 7.1

Topdressing (grams per container)

Container volume Low Medium High

1 qt (946 cm3) 2 4 6

1 gal (4,546 cm3) 6 17 26

5 gal (22,730 cm3) 26 70 105

Temperature

Longevity (months) oF oC

60 15 4 to 5

70 21 3 to 4

80 26 2 to 3

90 32 1 to 2

Table 12.6—Manufacturer’s recommendations for applying 

15-9-12 Osmocote® Plus controlled-release fertilizer. Adapted 

from Everris Company (2012).
needs to be handled carefully and not stored for long 

because much of the nutrients can be released because of 

prill breakage or high temperatures.

Growers can use the general recommendations provided 

by manufacturers if they classify their crops by relative nutrient uses: low, medium, or high (table 12.6). Of course, these 

applications rates should be used conservatively until their 

effect on individual plant growth and performance can be 

evaluated. Because CRF nutrient release is so affected by 

temperature, growers need to be cautious about using CRF 

label recommendations. Manufacturers base their release 

rates on an average temperature of about 70 oF (21 o

C), and 

those release rates increase by about 25 percent for every 9 o

F 

(5 o

C) increase in temperature (table 12.6). As with all nursery practices, growers should always initiate small scale trials 

before adopting any new fertilizer treatment.

Applying Organic Fertilizers

Composts could be incorporated into growing media 

but they must be fully mature to prevent fertilizer burn. 

One of the challenges of using liquid organic fertilizers is 

how to achieve the high soluble nitrogen levels necessary 

for rapid growth rates. High-quality nursery crops can be 

grown with organic fertilizers but, because their nutrient 

analysis is relatively low (figure 12.7B), production schedules may have to be adjusted.

Determining When to Fertilize

Because artificial growing media such as peat-vermiculite media are inherently infertile, fertilization should 

begin as soon as the seedlings or cuttings become established. Some brands of growing media contain a starter 

dose of fertilizer, however, so be sure and check the label. 

Homemade soil mixes that have been amended with 

compost or other organic fertilizers may not need fertilization right away but observe plant growth and establish 

small trials to be certain.

In nurseries, plant growth rates can be controlled by 

fertilization levels, especially nitrogen. As plants take up 

more nutrients, their growth rate increases rapidly until 

it reaches the critical point. After this point, adding more 

fertilizer does not increase plant growth but can be used 

246 Tropical Nursery Manual

Figure 12.11—Controlled-release fertilizers can be applied directly to containers (“topdressing”) if care is taken to achieve uniform 

application (A). Incorporating controlled-release fertilizers when the growing medium is mixed (B) helps to achieve even distribution of 

prills in small containers (C). Photo A by Brian F. Daley, photo B by Diane L. Haase, and photo C by Tara Luna.

B

A C
to “load” nursery stock with extra nutrients for use after 

outplanting. Beyond a certain point, however, excessive 

fertilization can cause a decline in plant growth and eventually results in toxicity (figure 12.12).

Some tropical plant species require very little fertilizer 

while others must be “pushed” with nitrogen to achieve 

good growth rates and reach target specifications. Smallseeded species expend their stored nutrients soon after 

germination whereas those with large seeds contain greater 

nutrient reserves and do not need to be fertilized right 

away. Excess fertilization early in the growing season can 

be detrimental to legumes and other natives, which must 

establish nitrogen-fixing bacteria or mycorrhizal fungi 

on their root systems before outplanting. Experience in 

growing a particular species is the best course of action to 

develop species-specific fertilizer prescriptions.

Tropical native plant growers should never wait for their 

crops to show deficiency symptoms before fertilizing. Plant 

growth rate will slow down before visible symptoms appear. 

Even after fertilization, it can take weeks before growth will 

resume following a deficient condition. Evaluating symptoms 

of nutrient deficiencies based on foliar characteristics can be 

challenging even for experts because different nutrient deficiencies have similar characteristic symptoms and considerable variation in these symptoms may occur among species. 

In addition, typical foliar symptoms, such as chlorosis, could 

be caused by something other than nutrient stress, such as 

heat damage or root disease. The position of the symptomatic 

foliage can be somewhat diagnostic. For instance, nitrogen is 

very mobile within the plant and will be translocated to new 

foliage when nitrogen is limiting. Therefore, nitrogen deficient plants show yellowing in the older rather than the newer 

foliage (figure 12.13A). Conversely, iron is very immobile in 

plants, so deficiency symptoms first appear in newer rather 

than older foliage (figure 12.13B). Keep in mind that excessive 

fertilization can cause toxicity symptoms (figure 12.13C). So, 

Plant Nutrition and Fertilization 247

Figure 12.12—Fertilization is one of 

the most effective ways to increase plant 

growth, which follows a characteristic 

pattern. As nutrient content increases 

in plant tissue, growth increases and 

deficiency symptoms are abated but 

fertilization beyond the optimum range 

is not always beneficial. Illustration by 

Jim Marin.

Figure 12.13—Nutrient deficiency symptoms such as chlorosis (yellowing) are common but can be caused by several different 

nutrients. Nitrogen chlorosis is seen first in older foliage (A), whereas iron chlorosis occurs in newer needles or leaves (B). Excessive 

fertilization can cause toxicity symptoms, such as necrosis along the leaf margin caused by boron toxicity (C). Photo A from Erdmann 

and others (1979), and photos B and C by Thomas D. Landis.

A B C
the take-home message is: foliar symptoms can be an indicator of nutrient (or other) problems but should never be used 

as the sole guide to fertilization. Crop monitoring and testing, 

experience, and knowledge about the growth phases are the 

best guides for determining fertilizer timing and rates.

Fertilization During Plant 

Growth Phases

Growers need to be aware of the different nutrient 

requirements during each growth phase and adjust fertilizer prescriptions accordingly (table 12.7). These adjustments are particularly important for nitrogen (especially the 

ammonium form of nitrogen), which tends to be a primary 

driver of plant growth and development (figure 12.14).

Establishment Phase

Fertilization needs to begin as soon as plants have 

emerged and have become established. Small plants are 

very succulent, however, and are particularly vulnerable 

to root damage from high salt concentrations. In addition, high nitrogen has been shown to increase the risk for 

damping-off fungi and other pest problems. Therefore, 

we recommend fertilizing with a low to moderate level of 

nitrogen (25 to 75 ppm) during this period.

Rapid Growth Phase

This phase is the period when plants attain most of 

their shoot development, and high levels of nitrogen (100 

to 150 ppm) tend to accelerate this growth. In addition to 

nitrogen, fertilizers need to include adequate levels of all 

the other mineral nutrients. Growers must closely monitor plant growth and development during the rapid growth 

phase, however, to ensure that shoots do not become excessively large or top heavy.

Hardening Phase

The objective of the hardening phase is to prepare plants 

for the stresses of shipping and outplanting by slowing shoot 

growth while simultaneously promoting stem and root 

growth. See Chapter 15, Hardening, for a complete description of this topic. High nitrogen levels stimulate plant growth, 

so reducing the amount of nitrogen (25 to 50 ppm) helps 

them to begin hardening. A lower ratio of nitrogen to that of 

phosphorus and potassium typically is helpful (table 12.7). In 

addition, changing to fertilizers containing the nitrate form 

of nitrogen (as opposed to ammonium) is helpful because 

nitrate-nitrogen does not promote shoot growth. Calcium 

nitrate is an ideal fertilizer for hardening because it provides 

the only soluble form of calcium, which has the added benefit of helping to promote strong cell wall development. It is 

important to distinguish granular calcium nitrate from liquid calcium ammonium nitrate, which does contain ammonium. A good rule of thumb for many tropical plants is that 

hardening should begin when shoots have reached 75 to 80 

percent of the target size. Plants take a few weeks to respond 

to this change in fertilization and will continue to grow after 

nitrogen fertilization has been reduced. Leaching the growing 

medium with several irrigations of plain water is a good way 

to make certain that all excess nitrogen is eliminated from the 

growing medium.

Monitoring and Testing

We recommend monitoring the electrical conductivity 

(EC) of fertilizer solutions and performing chemical analysis of plant foliage to determine if fertilization is sufficient, 

to prevent problems from developing, and to diagnose any 

deficiencies or toxicities if they do develop.

248 Tropical Nursery Manual

Growth phase Nitrogen Phosphorus Potassium

Establishment Medium High Low

Rapid growth High Medium Medium

Hardening Low Low High

Table 12.7—Examples of fertilization regimes adjusted for plant 

growth phases.

Figure 12.14—Because nitrogen is so critical to seedling physiology, nitrogen fertilization can be used to control the shoot-to-root 

ratio. It is also used to speed up or slow down plant growth. Fertilizers with the nitrate form of nitrogen are recommended during the 

hardening phase to slow shoot growth while stimulating roots; ammonium nitrogen is recommended during rapid growth. Illustration 

adapted from Dumroese and others (2008).
EC Testing

Because all fertilizers are taken up as electrically charged 

ions, the ability of a water solution to conduct electricity 

is an indication of how much fertilizer is present. Growers who fertigate need to periodically check the EC of the 

applied fertigation water and the growing medium solution. Measuring the EC of fertigation water as it is applied 

to the crop can confirm that the fertilizer solution has been 

correctly calculated and that the injector is functioning. 

Simple handheld EC meters (figure 12.15) are fairly 

inexpensive and are very useful for monitoring fertigation. 

The EC reading indicates the total amount of fertilizer salts 

and natural salts present in the water source. Normal readings in applied fertigation should range from 0.75 to 2.0 

uS/cm. Measuring the EC of water leached from containers 

can also help pinpoint problems of improper leaching and 

salt buildup within the growing medium. Measuring the 

EC of the solution in the growing medium, however, provides the best estimate of how much fertilizer is available to 

plant roots. The typical range of acceptable EC values in the 

growing medium for most tropical plant species is about 

1.2 to 2.5 uS/cm. If the EC is more than 2.5, it is a good idea 

to leach out the salts with clean irrigation water. See Chapter 11, Water Quality and Irrigation, for more information 

about EC measurements, guidelines, and units.

Foliar Testing

The best way to monitor plant nutrition and responses to 

fertilization is to test plant foliage for an exact measurement 

of nutrients that the plant has acquired. By examining tissue nutrient concentrations and simultaneously monitoring 

plant growth, it is possible to identify if and when specific 

nutrients are deficient or excessive/toxic (figure 12.12). Little 

published information is available for ideal mineral nutrient 

levels in tropical nursery plants so we recommend small trials to develop guidelines for your own species. Foliar samples must be collected in a systematic manner and sent to a 

reputable laboratory for processing (recommendations may 

be available through local extension agents). The analyzed 

nutrient concentration values are then compared with some 

known set of adequate nutrient values to determine which 

specific elements are deficient (table 12.8). The cost to analyze these samples is relatively inexpensive considering the 

potential improvement in crop quality that may result from 

having data to guide fertilizer regimes.

Growth Trials

Small growth trials are another good way to monitor 

plant nutrition and fertilization needs. These trials are 

especially informative for tropical plant species because 

so little published information is available. Detailed 

Plant Nutrition and Fertilization 249

Figure 12.15—An electrical conductivity meter used to estimate 

fertilizer salt concentrations in irrigation water, fertigation water, or 

saturated media extract. Photo by Douglass F. Jacobs.

Nutrient Range of foliar levels in 

healthy plants

Macronutrients (%)

Nitrogen 1.5 to 3.5

 Phosphorus 0.10 to 0.25

 Potassium 0.60 to 1.8

 Calcium 0.50 to 2.5

 Magnesium 0.15 to 0.50

 Sulfur 0.10 to 0.30

Micronutrients ppm 

(parts per million)

Iron 50 to 250

 Manganese 35 to 250

 Zinc 10 to 40

 Copper 5 to 20

 Boron 15 to 50

 Molybdenum 0.10 to 1.0

Table 12.8—Estimated ranges of foliar nutrient levels for healthy 

tropical plants (based on data compiled by Drechsel and Zech [1991] 

on field-grown, broad-leaved, tropical tree species). Nutrient ranges 

can vary greatly among species. Nursery trials are recommended to 

determine the best ranges for specific species.
documentation of growing conditions, fertilizer inputs, 

and resulting plant response can help to formulate future 

fertilizer prescriptions for a specific species within a nursery. See Chapter 20, Discovering Ways to Improve Nursery 

Practices and Plant Quality, for more information on how 

to make these discoveries through trials and experiments.

Reducing the Environmental 

Effects of Fertilization

Regardless of the method of fertilizer application or the 

type of fertilizer used, runoff of excess fertilizers is a major 

environmental concern. Nutrient ions, notably nitrate and 

phosphate, leach easily from container nurseries, and can pollute groundwater or adjacent streams. Many areas have laws 

and regulations limiting runoff and groundwater nitrate levels. Growers need to choose types of fertilizers and schedule 

their applications to minimize potential pollution concerns. 

Because nitrate and phosphate are so soluble in water, growers 

need to irrigate only when necessary and apply only enough 

water so that only small amounts drain out the bottom of the 

containers. This approach also makes sense from an economic 

standpoint, because the desire is to have most of the applied 

fertilizer taken up by crop plants rather than lost in runoff.

Reducing, eliminating, and managing environmental 

effects of fertilization in the nursery include these practices:

1. Apply fertilizer as part of a designed nutrient and irrigation program. The goal is never to push plants as quickly 

as possible through all phases of growth; instead, crops 

should be cultured to produce balanced, healthy plants 

for the best results after outplanting.

2. Consider using slow-release and controlled-release 

fertilizers in addition to, or instead of, highly soluble 

liquid fertilizers.

3. Select organic fertilizers that have less leaching potential.

4. Explore options to recycle or reuse irrigation water, 

including subirrigation systems or catchment ponds 

where runoff can be collected, treated, and reused.

5. Learn about water conservation and responsible 

management and reuse of runoff water, as discussed 

in Chapter 11, Water Quality and Irrigation.

A combination of these steps as appropriate for your 

nursery will help to minimize nutrient leaching and reduce 

the effects of fertilization on the environment.

References

Diver, S.; Greer, L.; Adam, K.L. 2008. Sustainable small-scale 

nursery production. Butte, MT: National Center for Appropriate Technology (NCAT) Sustainable Agriculture Project. https://

attra.ncat.org/attra-pub/summaries/summary.php?pub=60. 

(November 2011).

Drechsel, P.; Zech, W. 1991. Foliar nutrient levels of broad-leaved 

tropical trees: a tabular review. Plant and Soil. 131: 29–46.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Everris Company. 2012. Coated fertilizers. The Netherlands: 

Everris International, B.V. http://everris.us.com/plant-nutrition/

coated-fertilizers. (March 2012).

Landis, T.D.; Dumroese, R.K. 2011. Using organic fertilizers in forest and native plant nurseries. Forest Nursery Notes. 31(2): 9–18.

Morgenson, G. 1994. Using municipal organic wastes at LincolnOakes nurseries. In: Landis, T.D., comp. proceedings of the 

northeastern and intermountain forest and conservation nursery 

association meeting. Gen. Tech. Rep. RM-GTR-243. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky 

Mountain Forest and Range Experiment Station. 65–67.

 Additional Reading

Amaranthus, M.A. 2011. What are mycorrhizae? Grants Pass, 

OR: Mycorrhizal Applications Inc. http://www.mycorrhizae.

com/. (14 May 2011).

California Plant Health Association. 2002. Western fertilizer handbook. 9th ed. Danville, IL: Interstate Publishers. 356 p.

Card, A.; Whiting, D.; Wilson, C.; Reeder, J. 2009. Organic fertilizers. Fort Collins, CO: Colorado State University Extension, 

Colorado Master Gardener Program. CMG Garden Notes #234. 8 

p. http://www.cmg.colostate.edu. (May 2011).

Chaney, D.E.; Drinkwater, L.E.; Pettygrove, G.C. 1992. Organic 

soil amendments and fertilizers. Pub. No. 21505. Oakland, CA: 

University of California, Agriculture and Natural Resources. 35 p. 

Claassen, V.P.; Carey, J.L. 2007. Comparison of slow-release 

nitrogen yield from organic soil amendments and chemical fertilizers and implications for regeneration of disturbed sites. Land 

Degradation & Development. 18: 119–132.

Gaskell, M.; Smith, R. 2007. Nitrogen sources for organic vegetable crops. HortTechnology. 17: 431–441.

250 Tropical Nursery Manual
Hartz, T.K.; Smith, R.; Gaskell, M. 2010. Nitrogen availability 

from liquid organic fertilizers. HortTechnology. 20: 169–172.

Landis, T.D. 2011. Understanding and applying the carbon-tonitrogen ratio in nurseries. Forest Nursery Notes. 31(1): 10–15.

Landis, T.D.; Campbell, S.; Zensen, F. 1992. Agricultural pollution of surface water and groundwater in forest nurseries. In: 

Landis, T.D., tech. coord. Proceedings, Intermountain Forest 

Nursery Association, 1991. Gen. Tech. Rep. RM-GTR-211. Fort 

Collins, CO: U.S. Department of Agriculture, Forest Service, 

Rocky Mountain Forest and Range Experiment Station. 1–15.

Landis, T.D.; Khadduri, N. 2008. Composting applications in 

forest and conservation nurseries. Forest Nursery Notes. 28(2): 

9–18.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1989. 

The container tree nursery manual: volume 4, seedling nutrition 

and irrigation. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 119 p.

Meister Media. 2011. The Haber-Bosch Process. Washington, 

DC: Meister Media Worldwide. http://www.fertilizer101.org/. 

(8 November 2011).

Moral, R.; Paredes, C.; Bustamante, M.A.; Marhuenda-Egea, F.; 

Bernal, M.P. 2009. Utilisation of manure composts by high-value 

crops: safety and environmental challenges. Bioresource Technology. 100: 5454–5460.

The Scotts Company. 2006. Scotts fertilizer tech sheet. http://

www.scottsprohort.com/products/fertilizers/osmocote_plus.

cfm. (January 2006).

Rose, R.; Haase, D.L.; Boyer, D. 1995. Organic matter management in forest nurseries: theory and practice. Corvallis, OR: 

Oregon State University, Nursery Technology Cooperative. 65 p.

Roy, R.N.; Finck, A.; Blair, G.J.; Tandon, H.L.S. 2006. Plant nutrition for food security: a guide for integrated nutrient management. FAO Fertilizer and Plant Nutrition Bulletin 16. Rome, 

Italy: Food and Agriculture Organization of the United Nations. 

366 p. ftp://ftp.fao.org/docrep/fao/009/a0443e/a0443e.pdf. (April 

2011).

Sharpley, A.N.; Chapra, S.C.; Wedepohl, R.; Sims, J.T.; Daniels, 

T.C.; Reddy, K.R. 1994. Managing agricultural phosphorus for 

the protection of surface waters: issues and options. Journal of 

Environmental Quality. 23: 437–451.

Vaario, L.; Tervonen, A.; Haukioja, K.; Haukioja, M.; Pennanen, 

T.; Timonen, S. 2009. The effect of nursery substrate and fertilization on the growth and ectomycorrhizal status of containerized and outplanted seedlings of Picea abies. Canadian Journal of 

Forest Research. 39: 64–75.

Wikipedia. 2011a. Guano. http://en.wikipedia.org/wiki/Guano. 

(April 2011).

Wikipedia. 2011b. Sodium nitrate. http://en.wikipedia.org/wiki/

Sodium_nitrate. (April 2011).

Plant Nutrition and Fertilization 251
252
253

The web of life depends on microorganisms, a vast network of small, unseen 

allies that permeate the soil, water, and air of our planet. Many kinds of microorganisms existed for billions of years before any plants or animals came into 

being. Microorganisms created the atmosphere, turned bare rock and lava into 

soil, helped plants colonize land, and remain vital to the survival of plants, animals, and people today.

For people who work with plants, the greatest interest in microorganisms is in 

the complex living communities that are part of the soil. One gram (the weight of 

a small paperclip) of healthy soil can contain between 1 and 10 billion microorganisms. The living component of soil has a central role in ecosystem and plant 

health. Communities of bacteria, fungi, algae, protozoa, and other microorganisms make nutrients available to plants, create water and air channels, maintain 

soil structure, counterbalance pathogen populations, and recycle nutrients from 

organic matter that enable plants to grow.

This chapter focuses on two of the most important beneficial microorganisms 

for nurseries: nitrogen-fixing bacteria (rhizobia, the generic term for species in 

the genera Rhizobium and Bradyrhizobium)(figure 13.1) and mycorrhizal fungi 

(figure 13.2) that form mutually beneficial partnerships with their plant hosts. 

Scientists call this “mutualistic symbiosis.” In this manual, these beneficial microorganisms are called “microsymbionts.” Partnerships between beneficial microorganisms and plants are essential to plant health, as well as to healthy ecosystems 

and agro-ecosystems.

Beneficial Microorganisms

Kim M. Wilkinson and David P. Janos 13

Facing Page: The nitrogen-fixing bacteria rhizobia (Bradyrhizobium) form nodules on roots of legumes—in this case, on a 

native Acacia koa seedling in Hawai‘i. Photo by J.B. Friday.
In natural ecosystems, the root systems of most 

plants have microbial partnerships that enable them 

to survive and grow even in harsh conditions. In fact, 

microbial partnerships played a key role in enabling 

plants to first colonize land as they evolved out of the 

sea. Without microsymbiont partners, plants remain 

stunted and often die. These failures are frequently 

attributed to poor nursery stock when the real problem 

was the lack of the proper microsymbionts. In the nursery, microsymbionts can be introduced by “inoculating” 

the root systems of plants with the appropriate beneficial 

microorganisms to form effective partnerships.

254 Tropical Nursery Manual

Figure 13.2—“Myco” means “fungus” and “rhiza” means 

“root;” the word “mycorrhizae” means “fungus-roots.” Mycorrhizal root tip (arrow) on a pine tree root (the white filaments 

are the fungus extending beyond the root) (A). Dichotomously 

branched, ectomycorrhizal roots of Pinus elliottii in Florida; the 

finest rootlets are entirely ensheathed by white fungus filaments 

(B). Arbuscule of an arbuscular mycorrhizal fungus (C) that 

serves as a nutrient exchange site between the host plant and 

the fungus. Photo A by Thomas D. Landis, photo B by Tania 

Wyss, and photo C by Mark C. Brundrett.

Figure 13.1—Nitrogen-fixing bacteria include rhizobia, which 

form relationships with plants in the legume family. Pictured are 

native rhizobia nodules on the roots of the native Hawaiian forest 

tree koa (Acacia koa). Photo by J.B. Friday.

Mutualistic Symbiosis

Symbiosis technically refers to two or more organisms living intimately interconnected. As a scientific 

term, symbiosis can be mutualistic (both organisms 

benefit), parasitic (one organism benefits and the other 

is harmed), or commensal (one benefits, the other is 

unaffected). In popular usage, however, “symbiosis” 

is considered synonymous with “mutualistic symbiosis” 

—both organisms benefit. In this chapter, we employ 

the popular usage to refer to nitrogen-fixing bacteria 

and mycorrhizal fungi as “microsymbionts”—microorganisms that form a mutually beneficial partnership 

with their plant hosts.

B

C

A
The Importance of Beneficial 

Microorganisms in the Nursery

In natural ecosystems, the root systems of many plants 

have microbial partnerships with mycorrhizal fungi and, 

if applicable, with nitrogen-fixing bacteria. In the nursery, 

where plants have easy access to water and fertilizer, the 

benefits of these partnerships may not be apparent and 

their absence may go unnoticed. But in the field, plants 

need every advantage. Plants that have been inoculated 

in the nursery will be outplanted with microbial partnerships in place and often are better able to survive in the 

field. Noninoculated plants, however, must “fend for themselves” and establish microbial partnerships in the field. 

Many plantings take place on deforested or degraded land 

where native microsymbiont populations may be low or 

nonviable (figure 13.3).

Inoculating plants in the nursery is an opportunity to 

introduce select microsymbionts (figure 13.4). Similar to 

using seeds from specific seed sources, the nursery manager can match plants with optimal microsymbionts for 

specific site conditions. The presence of microsymbionts is 

often an important target plant characteristic.

Using microsymbionts in the nursery has the following 

benefits:

• Reduced environmental effects and fertilizer use in 

the nursery.

• Improved plant health and vigor.

• Improved resistance to disease.

• Better performance on outplanting sites.

Although this manual is for tropical nurseries, some tropical regions have montane habitats and species, and so a broad 

range of plant species and microsymbionts are mentioned in 

this chapter. Not all species and microsymbiont partners are 

present in a given region, so it is important to check about 

native species needs before using microsymbionts.

Beneficial Microorganisms 255

Figure 13.4—Plants with established microsymbiont partnerships often have a better chance of survival after outplanting. 

This photo shows a palm species, Bactris gasipaes; the plant on 

the right has a mycorrhizal partnership in place and the same 

age plant on the left does not. Photo by David P. Janos.

Figure 13.3—Microsymbionts often do not survive in the soil 

without their host plants. Native microsymbiont populations 

may be low on degraded sites, such as this formerly forested 

pasture in Hawai‘i (A) and this test bauxite surface mine in 

southwestern Costa Rica—when the aluminum-rich topsoil was 

removed from the mine, most mycorrhizal fungi were removed 

with it (B.) Photo A by J.B. Friday, and photo B by David P. Janos.

B

A
Nitrogen-Fixing Bacteria

Nitrogen is one of the most important nutrients for plant 

growth. Nitrogen (N2) is abundant in the Earth’s atmosphere, 

but the N2 gas must be converted to either nitrate (NO3

-

) or 

ammonium (NH4

+) ions before most plants can use it. In 

nature, nitrogen-fixing bacteria convert (“fix”) N2 from the 

air into a form usable to plants. When the growing roots of 

a plant capable of forming a partnership with rhizobia come 

in contact with a compatible strain of nitrogen-fixing bacteria in soil or growing media, the rhizobia bacteria will enter 

(“infect”) the roots. Nodules then form on the plant’s roots 

where the contact occurred. The bacteria live and multiply 

in the nodules on the host’s root system, providing nitrogen 

from the atmosphere to their plant host (figure 13.5). Each 

nodule contains millions of the bacteria that convert atmospheric nitrogen.

Although plants that form a partnership with nitrogenfixing bacteria are sometimes called “nitrogen-fixing plants” 

or “nitrogen-fixing trees,” the plant itself is unable to obtain 

atmospheric nitrogen. Through the mutualistic symbiotic 

partnership, the bacteria give nitrogen accumulated from 

the atmosphere to the plant, and in exchange, the bacteria 

get energy in the form of carbohydrates from the plant (Singleton and others 1990). When the host plant sheds leaves, 

256 Tropical Nursery Manual

Figure 13.5—The Nitrogen Cycle. All nitrogen in plants originates as an atmospheric gas, which is fixed by microorganisms (such 

as rhizobia and Frankia), fixed by humans in fertilizers by an energy-intensive industrial process, or to a very minor extent fixed 

by lightning or volcanism. The dashed lines in the diagram represent minor pathways; the solid lines represent major pathways. 

Adapted from Brown and Johnson (1996) by Jim Marin.
dies back, or dies, the nitrogen stored in the plant’s tissues 

is cycled throughout the ecosystem. The process of nitrogen 

fixation provides the major source of nitrogen fertility in 

tropical ecosystems (figure 13.5).

In the early 20th century, human beings also learned to 

convert atmospheric nitrogen gas into fertilizers through an 

energy-intensive industrial process called the Haber-Bosch 

process. The process requires a source of hydrogen gas to 

react with nitrogen from the air under heat and high pressure. The most common sources for the hydrogen are fossil 

fuels, and additional energy is expended to power the reaction. Growers who use synthetic nitrogen fertilizers such as 

ammonium nitrate and urea are using products generated by 

this process. In contrast to industrial/synthetic nitrogen fixation, biological nitrogen fixation by bacteria associated with 

green plants is powered by the sun, and thereby is renewable 

and effectively inexhaustible.

Nitrogen-fixing trees and plants are usually outplanted 

to help restore fertility, nutrient cycling, and organic matter 

to the ecosystem. Soils at the outplanting site, however, may 

not contain a viable strain of bacteria to form an effective 

partnership with the plant. Inoculating plants in the nursery 

ensures that an effective partnership is formed to enhance 

plant survival and growth and to accelerate rehabilitation of 

degraded land. Unlike mycorrhizal fungi, which affect most 

trees and plants, only a fraction of plants can form partnerships with nitrogen-fixing bacteria; however, nitrogen-fixing 

plants play a vital role in nutrient cycling.

Two types of nitrogen-fixing bacteria form symbiotic 

partnerships with plants: rhizobia (consisting of several genera) and the genus Frankia. Rhizobia nodulate many (but 

not all) members of the legume family (Fabaceae, sometimes called Leguminosae) (figure 13.6). The legume family is made up of three subfamilies (Mimosoideae, Caesalpinioideae, and Papilionoideae [Faboideae]). Rhizobia also 

nodulate species of the genus Parasponia in the elm family 

(Ulmaceae). Frankia are a genus of filamentous bacteria 

(in a group called “actinomycetes” because of their somewhat fungus-like appearance) that form partnerships with 

about 200 different plant species distributed across eight 

families (figure 13.7). The species affected by Frankia

are called “actinorhizal” plants (table 13.1).

Beneficial Microorganisms 257

Figure 13.6—Because nitrogen-fixing species improve soil fertility on degraded lands, they are widely used for restoration and 

sustainable agriculture. Examples of native Hawaiian legume species that form relationships with nitrogen-fixing rhizobia bacteria include Caesalpinia kavaiensis (Uhiuhi) (A) and Sophora chrysophylla (māmane)(B). Other nitrogen-fixing species are known 

throughout much of the tropics, such as Samanea (C), Gliricidia (D), and Sesbania (E) species. Some nitrogen-fixing trees are considered 

weeds outside their native range, such as Leucaena species (F). Photos A through E by J.B. Friday, and photo F by Tara Luna.

B

C E

F

A

D
Benefits of Inoculating With 

Nitrogen-Fixing Bacteria

Applications of inoculant for nitrogen-fixing bacteria 

can have some direct benefits in the nursery. If an effective 

partnership is formed, most of the plant’s requirements for 

nitrogen will be met, thereby reducing or eliminating the 

need to apply nitrogen fertilizer and decreasing the nursery’s need to manage pollution from fertilizer runoff.

In the field, however, is where the benefits of the 

partnership are most apparent. Nitrogen-fixing trees 

and plants sent from the nursery with their root systems 

already nodulated have faster early growth than plants that 

were not inoculated. Nursery inoculation can reduce costs 

in establishment and maintenance. The benefit from a few 

dollars’ worth of inoculant applied in the nursery not only 

offsets the need for purchased nitrogen fertilizer but is also 

much cheaper than replacing a tree that dies from nitrogen 

deficiency. Also, instead of providing spurts of fertilizers 

in the field (which may benefit surrounding weeds as well 

as the desired plant), the natural nitrogen fixation process 

provides a steady supply of nitrogen for the plant’s growth. 

Faster early field growth can lead to faster canopy closure, 

which in turn shades the soil and understory, reduces weed 

management expenses, and leads to faster restoration of 

the natural nutrient cycling and fertility role of nitrogenfixing species in the ecosystem.

Inoculating in the nursery ensures both the effectiveness 

and the timeliness of the nitrogen-fixing partnership. Noninoculated plants may eventually form a partnership in the 

field with a Frankia or rhizobia strain if some of the bacteria 

are present on the outplanting site. This partnership does 

not guarantee, however, that the plant will benefit. Some 

258 Tropical Nursery Manual

Bacteria Plant family Subfamily (notes) Examples (genus)

Rhizobia Legume (Fabaceae)

Caesalpinioideae

(about 1,900 species; about 23% fix nitrogen) Cassia and Senna

Mimosoideae

(about 2,800 species; about 90% fix nitrogen)

Enterolobium, Leucaena, 

Pithecellobium, Acacia, Albizia, 

Prosopis, and Mimosa

Papilionoideae

(about 12,300 species; about 97% fix nitrogen)

Sesbania, Cajanus, Erythrina, 

Gliricidia, and Robinia

Frankia

Birch (Betulaceae) Alnus

She-oak (Casuarinaceae) Casuarina, Allocasuarina, and

Gymnostoma

Coriariaceae Coriaria

Datiscaceae Datisca

Buckthorn (Rhamnaceae) Ceanothus and Rhamnus

Myrtle (Myricaceae) Myrica, Comptonia, and Myrtus

Oleaster (Elaeagnaceae) Elaeagnus and Hippophae

Rose (Rosaceae)

Cercocarpus, Chamaebatia, 

Cowania, Purshia, and

Chamaebatiaria

Table 13.1—Plants that form partnerships with nitrogen-fixing bacteria. Adapted from NFTA (1989) and Wall (2000).

Figure 13.7— Non-leguminous species that form relationships with 

nitrogen-fixing Frankia bacteria include Casuarina species. Photo by 

J.B. Friday.
plants will nodulate with a broad range of rhizobia strains, 

and not all strains are equally effective at fixing nitrogen 

(Keyser 2002). Some strains are very effective and productive, supplying plenty of nitrogen at a low cost to the host 

plant. Other strains are not productive, requiring a great deal 

of energy from the host plant with little return of nitrogen 

(figure 13.8). In other words, partnerships can range from 

mutually beneficial to parasitic (Evans 2002, Schmidt 2007, 

Baker and others 2009). Selecting microsymbionts that form 

healthy, productive partnerships is believed to warrant as 

much mindfulness as selecting seed sources (Schmidt 2007). 

In addition to source, time is also a factor for noninoculated 

plants after outplanting. It can take months or even years for 

effective partnerships to form if microsymbiont populations 

in the soil are low or inactive. Until the partnership forms, 

the plants are dependent on inputs of nitrogen fertilizers or 

the nitrogen available in the soil. Without fertilizer on poor 

sites, noninoculated plants will grow very slowly, and sometimes are outcompeted by weeds.

Acquiring Inoculants for 

Nitrogen-Fixing Bacteria

Inoculants are live nitrogen-fixing bacteria cultures 

that are applied to seeds or young plants, imparting the 

beneficial bacteria to the plant’s root system. Inoculants 

for nitrogen-fixing bacteria tend to be very specialized. In 

other words, they are not “one size fits all.” Care must be 

taken to select appropriate and effective nitrogen-fixing 

partners for specific plant species. Two forms of inoculant 

can be used for nitrogen-fixing plants in the nursery: pureculture inoculant is purchased from commercial suppliers, 

seed banks, or sometimes, universities (figure 13.9) and 

homemade (often called “crude”) inoculant is made from 

nodules collected from the roots of healthy nitrogen-fixing 

Beneficial Microorganisms 259

Figure 13.8—Some legumes can nodulate with a broad range of 

rhizobia strains, but not all strains are equally effective at fixing 

nitrogen. These tropical legume seedlings were inoculated with 

different rhizobia strains. Some partnerships were very effective 

(a green, thriving plant such as the one labeled 18b indicates that 

adequate nitrogen is being fixed at a low cost to the host plant) 

and others were not (a plant such as 4b is a little green, but not 

thriving and a plant such as 8b has no green). Careful selection 

of microsymbiont partners is important to ensure a productive 

partnership. Photo by Harold Keyser.

Figure 13.9—Nitrogen-fixing bacteria are commercially available as pure-culture inoculant (A), often in a carrier (B). Photo A 

by Tara Luna, and photo B by Mike Evans.

B

A
plants of the same species to be inoculated (figure 13.10). 

Whichever form is used, care should be taken when handling 

nitrogen-fixing bacteria inoculants because they are very perishable. These soil bacteria live underground in moist, dark 

conditions with relatively stable, cool temperatures. Similar 

conditions need to be maintained to ensure the viability of 

inoculant during storage, handling, and application.

Using Pure-Culture Inoculant

Pure-culture inoculants of nitrogen-fixing bacteria usually come in small packets of finely ground peat moss. Some 

manufactured inoculants contain select strains that have 

been tested for forming optimally productive partnerships 

with their host species. Select-strain inoculants should be 

considered if they can be obtained; these inoculants contain optimal partners for the host species to which they 

are matched, providing a good supply of nitrogen at a low 

cost to the plant. Superior strains can yield significant differences in the productivity and growth rate of the host 

plant; in some cases, they yield more than 40 percent better growth (Schmidt 2000). Not all manufactured inoculants are selected and matched to native species, however, 

so be sure to check the source. If a match cannot be found, 

use the crude inoculant method instead. Manufactured 

products usually come with application instructions; these 

directions need to be followed. In general, about 3.5 oz (100 

g) of cultured inoculant is sufficient to inoculate up to 3,000 

plants, usually exceeding the recommended 100,000 bacteria per plant. Because they contain living cultures of bacteria, these inoculants are perishable and need to be kept in 

cool, dark conditions, such as inside a refrigerator.

Peat-based inoculants are added to chlorine-free water to 

create a slurry. (If the nursery’s water supply is chlorinated, 

allowing a bucket of water to stand uncovered for 24 hours 

is a good way to let the chlorine evaporate.) A blender or 

electric mixer is recommended to blend the inoculant with 

water to ensure the bacteria are evenly mixed in the solution. 

If a blender is not available, a whisk can be used. After plants 

begin to nodulate, nodules from their roots can serve as the 

basis for making crude inoculant to use on future crops. This 

way, inoculant need be purchased only once for each plant 

species and can thereafter be perpetuated in the nursery.

Preparing Crude Inoculant

Nodules, the small root structures that house the bacteria, are used to make crude inoculant. Nodules can be col260 Tropical Nursery Manual

Figure 13.10—To make crude rhizobia 

inoculant, pick nodules off healthy plants 

in the nursery (A) or in the field. Make sure 

the nodules are active (indicated by a pink 

to red color inside)(B). Then, blend the nodules collected from root systems (C) in clean, 

chlorine-free water (D) and immediately water onto 2-week-old seedlings (E). Photos by 

Craig R. Elevitch, courtesy of Wilkinson and 

Elevitch 2003.

B

E

A C

D
lected from the roots of nursery stock that were previously 

inoculated with cultured, select inoculant, or nodules can 

be collected from healthy, established host plants. For rhizobia, a brown, pink, or red color inside is usually a good 

indicator that the millions of bacteria in the nodule are 

actively fixing nitrogen. For Frankia, desirable nodules will 

be white or yellow inside. Grey or green nodules should be 

avoided, because they likely are inactive.

To make crude inoculant, choose healthy, vigorous plants 

of the same species as the plants to be inoculated. Dig shallowly around the base of the nodulated plant to expose some 

of its root system. Young roots often contain the most active 

nodules. Search for nodules with the proper color and pick 

them off cleanly (figures 13.10A, 13.10B). Collect nodules 

from several healthy plants of the same species to ensure 

diversity (figure 13.10C). Put the nodules in a plastic bag 

or container and place them in a cooler for protection from 

direct sunlight and heat. As soon as possible after collection 

(within a few hours), put the nodules in a blender with clean, 

chlorine-free water (figure 13.10D). About 50 to 100 nodules 

blended in about 1 qt (1 L) of water are enough to inoculate 

about 500 plants. This solution is a homemade liquid inoculant, ready to apply in the same way as cultured inoculant 

(figure 13.10E).

Applying Inoculant

Inoculant for nitrogen-fixing bacteria is commonly 

applied when seedlings are emerging, usually within 2 

weeks of sowing, or just after cuttings have formed roots. 

This helps ensure successful nodulation and maximizes 

the benefits of using inoculants. The liquefied inoculant, 

made from either nodules or cultured inoculant as per the 

instructions in the previous sections, is then watered into 

the growing media or soil in which seedlings are growing 

(figure 13.10E).

Verifying the Nitrogen-Fixing Partnership

After 2 to 6 weeks, the noticeable signs in the following list 

should appear and are indications that the plant has formed 

a symbiotic partnership with nitrogen-fixing bacteria:

• Plants begin to grow well and are deep green despite 

the absence of added nitrogen fertilizer (figure 

13.11A).

• Root systems give off a faint but distinctive ammonialike scent.

• Nodules are visible on the root system.

• When a nodule is broken open, its inside is pink, red, 

or brown (for rhizobia) (figure 13.11B), or yellow or 

white (for Frankia).

Management Considerations

As with any nursery practice, becoming familiar 

with the application and management of nitrogen-fixing 

microsymbionts is a learning process. Several factors are 

of primary concern to the nursery manager when using 

inoculants for nitrogen-fixing bacteria:

• Timing—Ensure the inoculant is applied when 

seedlings have just emerged or when cuttings have 

formed new roots to ensure successful nodulation and 

maximize the benefits of using inoculants.

• Fertilization and Micronutrients—The use of nitrogen-fixing bacterial inoculant requires some adjustments in fertilization. Excessive nitrogen fertilizer will 

inhibit formation of the partnership. If an optimal 

partnership is formed, the application of nitrogen 

may be eliminated from nitrogen-fixing plants and 

they may need to be isolated from nonnitrogen-fixing 

species to implement this change in fertilization. 

Beneficial Microorganisms 261

Figure 13.11—The 6-week-old native Acacia koa seedlings 

(right) were inoculated with rhizobia at 2 weeks of age; the 

seedlings on the left were not inoculated (A). Nodules from an 

Acacia koa seedling showing pink inside, signifying nitrogen is 

being fixed (B). Photo A by Craig R. Elevitch, and photo B by 

J.B. Friday.

B

A
Some nutrients, including calcium, potassium, molybdenum, and iron, are necessary to facilitate nodulation. These nutrients need to be incorporated into the 

growing medium. Phosphorus is also necessary for 

nodulation, supplied from the growing medium or, 

better yet, through mycorrhizal partnerships.

• Water Quality—Excessive chlorine in water is detrimental to rhizobia and Frankia. The water supply 

may need to be tested and a chlorine filter obtained 

if excessive chlorine is a problem in the water supply. As an alternative, chlorine will evaporate if clean 

water is left to stand uncovered in a container for 24 

hours before use.

• Sourcing Inoculants—Locating appropriate sources 

of viable inoculants (either cultured or obtained 

as nodules) matched to native species may require 

some research and time but benefits of successful 

inoculation are well worth the effort.

• Client Education—Make sure nursery clients 

and people outplanting the plants understand the 

nitrogen-fixing bacteria so they appreciate the presence of nodules and are careful not to expose root 

systems to full sun. Also, educate clients so they 

know that only some species of plants can form this 

partnership—otherwise some might think all species 

can fix nitrogen.

• Outplanting Site Considerations—After nodules 

form, rhizobia are enclosed and usually less affected 

by soil conditions such as pH or aluminum toxicity 

than are other beneficial microorganisms such as 

mycorrhizal fungi. In very harsh outplanting conditions (such as extremely low pH in some rehabilitation sites), however, the rhizobia are not likely to 

spread in the soil, and so might not be available 

to spread to subsequent cohorts of the outplanted 

species. Therefore, inoculating subsequent crops in 

the nursery before outplanting on the same site is 

advisable.

Tripartite Symbiosis

Most plants that form partnerships with nitrogen-fixing 

bacteria also require mycorrhizal partners. When a nitrogenfixing plant has effective partnerships with both nitrogenfixing bacteria and mycorrhizal fungi, this is called “tripartite symbiosis” because three partners exist (a host plant and 

two microsymbionts) (figure 13.12). When working with 

both types of microsymbionts, simply apply each inoculant 

separately, as described in the sections of this chapter.

Several studies have shown that legumes may need to 

first form arbuscular mycorrhizae before they can form 

rhizobia nodules. This may occur because nitrogen fixation 

is an energy-demanding process, and the plant’s energy 

metabolism is highly phosphorus-dependent. In other 

words, a grower who applies only rhizobia inoculant may 

have difficulty getting good nodulation if the mycorrhizal 

partnership is not in place (or if adequate phosphorus is 

not available in the growing medium). The next section 

describes how to introduce mycorrhizal fungi to nursery 

crops—they often are incorporated in the growing medium 

before the rhizobia inoculant is applied.

Mycorrhizal Fungi

Unlike nitrogen-fixing bacteria, mycorrhizal fungi form 

partnerships with nearly all plant families and forest trees. 

“Myco” means “fungus” and “rhiza” means “root;” the 

word “mycorrhizae” means “fungus-roots.” Most of the 

world’s plants depend on their partnership with mycorrhizal fungi to grow and thrive. The host plant’s roots provide 

a substrate for the fungi and supply food in the form of 

simple carbohydrates. In exchange, the mycorrhizal fungi 

offer the following benefits to the host plant:

• Increased Water and Nutrient Uptake—Mycorrhizal 

fungi help plants absorb mineral nutrients, especially 

nitrogen, phosphorus, and several micronutrients 

such as zinc and copper. The fungal hyphae extend 

out into the soil far beyond the host’s roots, expanding the mineral- and water-absorbing surface area for 

the host plant. Researchers estimate that mycorrhizal 

fungus hyphae can explore volumes of soil hundreds 

to thousands of times greater than roots can alone.

• Stress and Disease Protection—Mycorrhizal fungi 

protect the plant host in several ways. With ectomycorrhizal fungi, for example, a fungus sheath (called a 

“mantle”) completely covers fragile root tips and acts 

262 Tropical Nursery Manual

Figure 13.12—The Gliricidia sepium plants on the far left were 

inoculated with both rhizobia (R) and mycorrhizal fungi (M) and 

show tripartite symbiosis: a beneficial partnership among the plant 

host, rhizobia bacteria, and arbuscular mycorrhizal fungi. Photo by 

Kenneth W. Mudge
as a physical barrier to dryness, pests, and toxic soil 

contaminants. Other mycorrhizal fungi may produce 

antibiotics, which provide chemical protection.

• Increased Vigor and Growth—Plants with mycorrhizal roots may have an improved hormone status, 

and they survive and grow better than noninoculated 

plants after they are planted out on a project site. 

Studies show that establishing a partnership with 

mycorrhizal fungi while the plants are in the nursery 

results in improved field growth (Habte and others 

2001, Baker and others 2009).

The following three types of mycorrhizae are important 

to tropical native plant nurseries (table 13.2):

• Arbuscular Mycorrhizae (AM)—Formed by the 

most ancient and predominant type of mycorrhizal fungi. AM fungi are found on the roots of most 

tropical plants and many of the world’s food crops 

(including rice, corn, and legumes), associating with 

more than 80 percent of the world’s plant families 

(figure 13.13A).

• Ectomycorrhizae (ECM)—Partnerships with many 

temperate forest trees and a few abundant tropical 

trees including pines (Pinus), eucalypts (Eucalyptus), poplars (Populus), and dipterocarps (Dipterocarpus) (figure 13.13B).

• Ericoid Mycorrhizae (ERM)—Partnerships with 

plants in the heath or heather (Ericaceae) family, 

including the genera of blueberries, cranberries, 

azaleas, and rhododendrons (figure 13.13C).

Mycorrhizal fungi are not “one size fits all,” but they 

often are “one size fits many.” Also, one plant can partner simultaneously with several species of mycorrhizal 

fungi, and a plant may change partners over time as it 

grows and adapts to its environment (Amaranthus 2010). 

Beneficial Microorganisms 263

Mycorrhizal fungi Plants

Arbuscular 

mycorrhizal (AM) 

More than 80% of the world’s plant families 

including most tropical trees, herbs, and 

ferns

Ectomycorrhizal 

(ECM) 

Fewer than 10% of plant families, 

including the genera pine (Pinus), oak 

(Quercus), eucalyptus (Eucalyptus)

AM and ECM

Some Allocasuarina, Acacia, Eucalyptus, 

juniper (Juniperus), poplar (Populus), and 

willow (Salix)

Ericoid mycorrhizal 

(ERM)

Heather or heath family (Ericaceae in 

the broad sense, including the former 

Epacridaceae and Empetraceae), including 

blueberry (Vaccinium) and Rhododendron

(including azaleas)

Table 13.2—Plants and their mycorrhizal partners. Adapted 

from Castellano and Molina (1990) and Wang and Qiu (2006).

Figure 13.13—The three types of mycorrhizal fungi. Arbuscular mycorrhizal (AM) fungi (A). Ectomycorrhizal (ECM) fungi (B)—the 

mantles of ECM may be visible to the unaided eye, although ECM on Eucalyptus can be inconspicuous. Ericoid mycorrhizal (ERM) fungi (C). 

Both AM and ERM can only be seen on plant roots with the aid of a microscope. Photos A and B by Michael A. Castellano, and photo C by 

Efren Cazares.

A B C
Because most plants are associated with a particular type 

of mycorrhizal fungus, however, different plant species 

have different fungal partners that must be matched 

appropriately to be effective (table 13.2).

Mycorrhizal fungi may be obtained either from commercial suppliers or from roots around a healthy host plant 

of the species being propagated. In all cases, mycorrhizal 

inoculum must physically contact living roots of the plant 

to colonize most effectively. Ways to acquire and successfully apply mycorrhizal fungi are explained in subsequent 

sections. Although the fungi are similar in how they function and in their benefits to host plants, they appear differently on roots. Also, each mycorrhizal fungi type has a 

unique application method that must be described separately. Management practices in the nursery are similar 

and will be discussed together at the end of this section.

Arbuscular Mycorrhizal (AM) Fungi

AM fungi are essential for most tropical trees and other 

plants and for many annual crops and grasses. AM fungi 

are not visible on plant roots to the unaided eye and must 

be observed under a microscope. The large spores of AM 

fungi are not easily disseminated by wind, unlike the winddispersed microscopic spores of ECM fungi.

Inoculant for AM fungi is sometimes collected from root 

systems of AM host plants or soil underneath them and 

incorporated into growing media. This method can work 

well because the fungi collected are likely to be adapted to 

prevailing site conditions (Janos and others 2001). Freshly 

collected and chopped mycorrhizal roots must be used 

within six days of collection or their efficacy will decline. 

This method is often discouraged, however, because of 

damage to plants and natural ecosystems, variable effectiveness, and the risk of introducing pests and pathogens 

along with the soil or roots. The two main sources of AM 

fungi inoculant for nurseries are “pot culture” made from 

a known fungus species, and commercially available cultures. Because AM spores are relatively large, it is critical to 

ensure that spores come in direct contact with the root systems. Spores will not pass easily through irrigation injectors or nozzles. Therefore, for all inoculant types, thorough 

incorporation into growing media is the best practice.

Pot Culture Inoculant

In pot culture inoculant, a specific AM fungus species 

is acquired either commercially or from a field site as a 

starter culture and then incorporated into a sterile growing medium. A host plant such as corn, sorghum, clover, or 

an herbaceous native plant, is grown in this substrate. As 

the host plant grows, the AM fungi multiply in the medium 

(figure 13.14). After the host plant roots have spread 

throughout the medium, their shoots are removed and 

the substrate, now rich in roots, spores, and mycelium, is 

chopped up and incorporated into fresh growing medium 

before containers are filled and seeds are sown or cuttings 

stuck. This technique is highly effective for propagating 

AM fungi in the nursery. For further details on how to use 

this method, consult the publications in the References section of this chapter (particularly Habte and Osorio 2001, 

Miyasaka and others 2003).

Commercial Inoculant Sources

Commercial sources of AM fungi inoculant are also 

available, usually containing several species or strains. 

Because AM fungus spores are fragile, they are usually 

mixed with a carrier such as vermiculite or calcined clay 

to aid in application. These products are thoroughly incorporated into the growing medium before filling containers. 

264 Tropical Nursery Manual

Figure 13.14—In pot culture inoculant, a specific arbuscular 

mycorrhizal fungus species is acquired as a starter culture and added to a sterile growing medium with a fast-growing host plant (A). 

The shoots of host plants are later removed, and the substrate, now 

rich in roots, spores, and mycelium, is chopped up (B) and incorporated into the growing medium before containers are filled. Photos 

by Thomas D. Landis.

B

A
Inoculation effectiveness has been shown to differ considerably between different products so it is wise to test before 

purchasing large quantities of a specific product. Laboratories can provide a live spore count per volume, which is the 

best measure of inoculum vigor.

Verifying the Effectiveness of AM Fungi Inoculation

To verify the effectiveness of AM fungi inoculation, 

roots must be stained and examined under a microscope 

(figure 13.15A). This verification can often be done through 

a soil scientist at a local agricultural extension office. After 

some practice, nursery staff may get a feel for when inoculation is successful, because noninoculated plants often 

grow more slowly and may have higher incidence of root 

rot issues. The plants also may exhibit signs of phosphorus 

deficiency (a frequent consequence of lack of mycorrhiza), 

indicated by purple coloration of leaves or other symptoms 

(figure 13.15B).

Ectomycorrhizal (ECM) Fungi

Many recognizable mushrooms are fruiting bodies of 

ECM fungi. These fruiting bodies are a small portion of the 

total organism; underground, the amount of fungus covering the short feeder roots of plants may be enormous. 

ECM fungi extend the volume of the feeding area of roots 

by many times and protectively coat the feeder roots. On 

some species, such as pines (Pinus), the mantles of ECM are 

visible on the roots of the host plant. For other species, such 

as many Eucalyptus ECM, the mantles can be inconspicuous. ECM are important to many temperate forest species, 

especially evergreens. In the tropics, far fewer plant species 

partner with ECM fungi than with AM fungi. In Hawai‘i, 

for example, no native ECM are known, although some 

strains may have been introduced along with introduced 

trees (Amaranthus 2010). ECM only affect a small percentage of tropical species, including pines, eucalypts, poplars, 

oaks, dipterocarps, and some legumes (table 13.3).

Four sources of ECM fungi inoculant have been used in 

nurseries. Nurse plants and soil spores have been used historically while spores and pure culture inoculant are usually recommended for nurseries.

Nurse Plants as Inoculum Sources

In the early days of trying to establish pines in the 

tropics where they were not native, “nurse” plants were 

sometimes used. Conspicuously vigorous mycorrhizal 

seedlings were transplanted to nursery beds at 3- to 6-ft 

(1- to 2-m) intervals and allowed to become established, 

maintaining the ECM fungi on their roots. Seeds were 

then sown or germinants transplanted around and between 

these mycorrhizal nurse plants. After becoming colonized 

by ECM fungi spreading from the nurse plants, the seedlings were transplanted to the outplanting site. Some plants 

were left in the beds to serve as nurse plants for the next 

crop of seedlings. Sometimes the spread of mycorrhizal 

fungi from the nurse seedlings was slow (roughly 2 ft per 

year), which suggested that some unfavorable soil property 

(such as high pH) should have been remedied. The mycorrhizal fungi usually spread fastest among seedlings in sterilized soil. This method was laborious, and, as with all bareroot crops, care had to be taken not to cause damage when 

lifting seedlings (Mikola 1973).

Beneficial Microorganisms 265

Figure 13.15—In this microscope photo (400 times magnification), a tropical tree root has been clarified and stained with a blue 

dye so that an AM fungus “vesicle” within the walls of a root cortical cell is clearly visible (A). Vesicles are places where AM fungi 

store excess energy-rich materials, such as lipids. Vesicles always are attached to AM fungus filament (a hypha), which distinguishes 

them from the spore-bearing structures of some root-parasitic fungi. Although AM fungi are visible only under a microscope, nursery 

workers may observe differences in plant growth of inoculated versus noninoculated plants. A mosaic pattern of nutrient deficiencies 

as shown by these mahogany (Swietenia species) seedlings, may indicate that some plants have formed successful partnerships while 

others have not (B). Photo A by David P. Janos, and photo B by Tara Luna.

A B
Soils as Inoculum Sources

Topsoil, humus, or duff from beneath ECM fungi host 

trees has historically been used to inoculate nursery plants. 

This practice is more common in bareroot nurseries in 

temperate regions than in container nurseries. Because 

sterilization would kill these beneficial fungi, unsterilized 

soil and organic matter are incorporated into the growing 

medium, up to 10 percent by volume. Today, this practice 

is generally discouraged for ECM fungi because (1) large 

quantities of soil are required, which can make the process 

labor intensive and have a detrimental effect on the natural 

ecosystem, (2) the quality and quantity of spores may be 

highly variable, and (3) pathogens may be introduced along 

with the inoculant. If soil is used, inoculum should be collected from plant communities near the outplanting site. 

Small amounts should be collected from several different 

sites, then thoroughly mixed, and care should be taken not 

to damage the plants during soil collection.

Spores as Inoculum Sources

Nurseries can make their own ECM inoculum from 

spores. Collected from the fruiting bodies (figure 13.16) of 

mushrooms, puffballs, and especially truffles, these fruiting bodies, full of spores, are rinsed, sliced, and pulverized 

in a blender for several minutes. The resulting thick liquid 

is diluted with water and poured into the growing media of 

germinating seedlings or newly rooted cuttings. Plants are 

usually inoculated 6 to 12 weeks after sowing (figure 13.17). 

Two applications 2 to 3 weeks apart are recommended to 

ensure even inoculation.

Pure-Culture Inoculum

ECM fungi are available commercially as pure cultures, 

usually in a peat-based carrier (figure 13.18). The quality 

of commercial sources varies, however, so it is important 

to verify vigor by testing formation of mycorrhizae. Most 

commercial sources contain several different species of 

266 Tropical Nursery Manual

Family Genera

Gnetaceae Gnetum

Pinaceae Cedrus, Keteleeria, Larix, Picea, and Pinus

Nyctaginaceae Guapira, Neea, and Pisonia 

Polygonaceae Coccoloba 

Myrtaceae Allosyncarpia, Agonis, Angophora, Baeckea, Eucalyptus, Leptospermum, Melaleuca, Tristania, and Tristaniopsis

Fabaceae: Caesalpinioideae 

Afzelia, Anthonotha, Aphanocalyx, Berlinia, Brachystegia, Cryptosepalum, Dicymbe, 

Didelotia, Eperua, Gilbertiodendron, Gleditsia, Intsia, Isoberlinia, Julbernardia, 

Microberlinia, Monopetalanthus, Paraberlinia, Paramacrolobium, Pellegriniodendron, 

Tetraberlinia, and Toubaouate 

Fabaceae: Papilionoideae Aldinia, Gastrolobium, Gompholobium, Jacksonia, Lonchocarpus, Mirbelia, Oxylobium, and Pericopsis 

Fabaceae: Mimosoideae Acacia and Calliandra 

Casuarinaceae Allocasuarina and Casuarina 

Fagaceae Castanea, Castanopsis, Fagus, Lithocarpus, and Quercus

Phyllanthaceae (Euphorbiaceae) Uapaca and Poranthera

Salicaceae Populus and Salix 

Rhamnaceae Cryptandra, Pomaderris, Spyridium, and Trymalium

Dipterocarpaceae Anisoptera, Dipterocarpus, Hopea, Marquesia, Monotes, Shorea, Vateria, Vateriopsis, and Vatica 

Sarcolaenaceae Leptolaena, Sarcolaena, and Schizolaena

Table 13.3—Tropical families and genera with ectomycorrhizal associations. Adapted from Brundrett (2009).
ECM fungi. Commercial inoculum can be purchased separately and mixed into the growing medium as per the 

instructions on the product and before filling containers. 

In some areas, bales of growing medium with inoculum 

already premixed may be purchased. It is important to 

inquire if selected strains to match site needs are available 

through suppliers.

Verifying the Effectiveness of ECM 

Fungi Inoculation

With practice, nursery staff can learn to recognize ECM 

fungi on the root systems of plants—they are fairly easy to 

see and often involve conspicuous morphological changes 

of the finest roots. During the hardening phase, short feeder 

roots need to be examined for a cottony white appearance 

on the root surface or a white or brightly colored mantle 

or sheath over the roots (figure 13.19A). Unlike pathogenic 

fungi, mycorrhizae never show signs of root decay. Sometimes, mushrooms or other fruiting bodies will appear 

in containers alongside their host plants (figure 13.19B). 

Although these structures are visible to the unaided eye, it 

is also recommended to send plant samples to a laboratory 

for verification. A local soil extension agent or university 

likely can assist with this process.

Beneficial Microorganisms 267

Figure 13.16—Fruiting bodies of ectomycorrhizal fungi. A puffball of Pisolithus 

tinctorius, showing the cavities where the 

spores are located (A). An Amanita from 

pine forest in Guatemala (B), and gilled 

ectomycorrhizal mushrooms and roots 

from dipterocarp forest in Malaysia (C). 

Photo A by Michelle M. Cram, and photos 

B and C by David P. Janos.

Figure 13.17—Inoculating tree seedlings with ectomycorrhizal fungi. Two applications 2 to 3 weeks apart are recommended 

to ensure even inoculation. Photo by Michael A. Castellano.

Figure 13.18—Forms of commercial ectomycorrhizal inoculants 

available for nurseries. Photo by Thomas D. Landis.

B

C

A
Ericoid Mycorrhizal (ERM) Fungi

Plants that form partnerships with ERM fungi are able to 

grow in exceptionally nitrogen-poor soils and harsh conditions, including bogs, alpine meadows, tundra, and even in 

soils with high concentrations of certain toxic metals (figure 13.20). ERM fungi form partnerships in the plant order 

Ericales in the heath (Epacridaceae), crowberry (Empetraceae), and most of the rhododendron (Ericaceae) family 

(table 13.4). Similar to ECM fungi and AM fungi, ERM fungi 

must come in contact with the host plants roots to form 

partnerships. Ericoid mycorrhizal inoculant is available as 

commercial cultures or from soil near healthy host plants. 

The product or soil is mixed into nursery growing medium. 

The fungus forms a net over the narrow “hair roots” (the 

fine, ultimate rootlets of the plants that are only a few cells 

268 Tropical Nursery Manual

Figure 13.19—Nursery staff can learn to recognize the presence 

or absence of ectomycorrhizal fungi by examining plants. Cottony 

white ectomycorrhizae may be visible on roots of some species (A), 

fruiting bodies may be growing from containers (B). Photo A by 

William Sayward, and photo B by Michael A. Castellano.

Figure 13.20—A native Hawaiian Ericoid, Vaccinium reticulatum, ‘ōhelo, growing on a recent lava flow. Partnerships 

with ericoid mycorrhizal fungi enable these plants to survive 

and thrive in harsh conditions. Photo by Kim M. Wilkinson.

Family Genera

Ericaceae

Acrotriche, Andersonia, Astroloma, Brachyloma, 

Cassiope, Calluna, Ceratiola, Conostephium, Corema, 

Cyathodes, Dracophyllum, Empetrum, Epacris, Erica, 

Gaultheria, Kalmia, Ledum, Leucopogon, Lissanthe, 

Lysinema, Melichrus, Monotoca, Needhamiella, 

Oligarrhena, 

Pentachondra, Richea, Rhododendron, Rupicola, 

Sphenotoma, Sprengelia, Styphelia, Trochocarpa, Vaccinium, Woollsia

Table 13.4—Genera known to associate with ericoid mycorrhizal 

fungi. Adapted from Read (1996) and Smith and Read (1997).

B

A
wide), infecting the outer cells. As with AM fungi, nutrients 

are shared through the membranes that form the boundary 

between the fungus and plant roots. Laboratory confirmation is recommended to verify that successful inoculation 

has taken place.

Management Considerations for 

Mycorrhizal Fungi

When using mycorrhizal inoculants for the first time, 

it is recommended to start small and evaluate a few techniques and sources. Compare some trays or benches with 

and without mycorrhizae to determine how management 

and scheduling need to be modified to culture mycorrhizal 

roots. In some cases, working with a manufactured product 

of known quality may be the easiest way to begin; the nursery can then expand into collecting and processing its own 

inoculant sources. Monitor the effectiveness of inoculation 

and keep records of crop development. See Chapter 20, 

Discovering Ways to Improve Nursery Practices and Plant 

Quality, to learn more about how to create some small trials 

and experiments.

Although mycorrhizal fungi are not very specialized, 

different strains of mycorrhizae are believed to perform 

differently for given site challenges. Some select or pureculture inoculants may support high productivity in certain site conditions but may be less productive than native 

strains on other sites. For example, some strains may be 

more beneficial if lack of nutrients is the main challenge 

while others may be particularly helpful to their hosts in 

withstanding soil pathogens or even heavy metals. If possible, working with several strains for diversity in the 

field may be a good safeguard, especially because plants 

can partner with multiple strains simultaneously and can 

change partners if necessary to adapt to site conditions. 

The nursery can do a little research or work with a specialist to help with the following tasks:

• Select optimal mycorrhizal partners for the species 

and outplanting sites.

• Determine the most appropriate sources of inoculant 

and evaluate their effectiveness in the nursery.

• Design outplanting trials to evaluate plant vigor and 

survival and modify the inoculant sources if improvements are needed.

Beneficial Microorganisms 269

Figure 13.21—Four-month-old guava seedlings (Psidium guajava) in the nursery in a low fertility (about 8 ppm available phosphorus) lowland tropical acid clay soil; the plant on the right has a mycorrhizal partnership in place while the plant on the left does not (A). 

Even with abundant phosphorus fertilization, lack of mycorrhizal fungi can slow plant growth. The photo shows lychee (Litchi chinensis) 

air layers grown for 16 months in 25 gal pots of a soil-free medium after cutting from the source trees (B). Although phosphorus fertilization 

did not affect growth, AM fungus inoculation with field-collected inoculant improved shoot growth by 39 percent (see Janos and others 

2001). Photos by David P. Janos.

B

A
Inoculation with mycorrhizal fungi affects plant growth 

(figure 13.21). Fertilization practices will need to be adjusted 

to support the formation of mycorrhizal partnerships in the 

nursery. An excessive amount of phosphorus inhibits formation of the partnership; therefore phosphorus must be 

reduced. Enough phosphorus must be present to keep the 

mycorrhizal fungi from competing with its host plant for 

the nutrient, however, and potentially becoming parasitic 

instead of symbiotic. The recommendation is to use “low 

but sufficient” levels of phosphorus to facilitate the partnership (Miyasaka and others 2003). In some cases, the overall 

quantity of fertilizer may need to be reduced by half or more 

because of the efficiency of nutrient uptake by mycorrhizal 

fungi. Fertilizer type and form is also important. If nitrogen is applied, ammonium-nitrate is better used by a wide 

variety of plants than nitrate-nitrogen alone (Castellano and 

Molina 1990). In general, controlled-release fertilizers may 

be better than liquid fertilizers for inoculated plants because 

they release small doses of nutrients gradually rather than 

sudden high doses periodically. Excessive or inadequate 

water will inhibit the presence of mycorrhizal fungi and the 

formation of the partnership, so watering schedules need to 

be modified accordingly. Nursery staff who are willing to be 

observant and flexible as the nursery embarks on the use of 

mycorrhizal fungi will be the best decision makers in terms 

of modifying fertilization and watering regimes to support 

the microsymbionts.

Other management adjustments may be necessary 

because of improved survival and growth. Improved 

survival percentages will affect estimates and oversow 

rates. Scheduling also may be affected; inoculated plants 

may be ready for outplanting sooner than noninoculated 

plants. Applications of certain fungicides are detrimental 

to mycorrhizal fungi; susceptibility varies by species and 

pesticide applications need to be assessed and adjusted.

Some plants form partnerships with both AM fungi and 

ECM fungi. These plants include some of the Allocasuarina, 

Acacia, Eucalyptus, Juniperus, and Populus species. In these 

cases, trials should be done to see which inoculant produces 

the best results in the nursery, and if those results persist 

after outplanting. For some species, additional inoculation 

with the second type of mycorrhiza (usually ECM fungi following AM fungi) may be necessary before outplanting.

Also, most plants that form partnerships with nitrogen-fixing bacteria also require mycorrhizal partners. 

For example, many leguminous trees partner with both rhizobia and AM fungi; in these cases, it may be beneficial to 

apply the AM fungi before the rhizobia (as discussed previously). Trees such as alders (Alnus) partner with both Frankia and ECM fungi. In these cases, inoculants can be applied 

separately to the same crop of plants.

Other Beneficial Microorganisms

In natural soil, communities of bacteria, fungi, algae, 

protozoa, and other microorganisms make nutrients available to plants, create channels for water and air, maintain 

soil structure, and cycle nutrients and organic matter. A 

healthy population of soil microorganisms helps to maintain ecological balance, preventing the onset of major problems from soil viruses or other pathogens. Realizing that 

soil is alive, and reducing or eliminating practices that may 

be harmful to soil microlife is important. As Aldo Leopold 

advised, “The first rule of intelligent tinkering is to keep 

all the parts.” Using compost, mulch, and organic matter 

is important for soil life. Protecting soil from erosion and 

unnecessary disturbances, eliminating pollution from soluble fertilizers, and minimizing the use of fungicides, disinfectants, and other chemicals that may kill microlife are 

all key practices. Introductions of mycorrhizal fungi and 

nitrogen-fixing bacteria in the nursery can support the balance of beneficial soil microorganisms.

 Acknowledgements

The authors thank the following people for sharing their assistance and expertise while this chapter was being developed: 

Mike Amaranthus, Grants Pass, OR. Microbiologist; Adjunct Associate Professor, Oregon State University; President, Mycorrhizal 

Applications, Inc.

Mitiku Habte, Honolulu, HI. Professor of Soil Science, University of 

Hawai‘i at Mānoa Department of Tropical Plant and Soil Sciences.

Harold Keyser, Kahului, HI. Maui County Administrator, University of Hawai‘i College of Tropical Agriculture and Human 

Resources (CTAHR). 

Jim Trappe, Corvallis, OR. Professor, Oregon State University, 

Department of Forest Science. 

Kenneth Mudge, Ithaca, NY. Associate Professor, Cornell University Department of Horticulture.

References

Amaranthus, M. 2010. Personal communication. Grant’s Pass, 

OR. Microbiologist; Adjunct Associate Professor, Oregon State 

University; President, Mycorrhizal Applications, Inc. 

Baker, P.J.; Scowcroft, P.G.; Ewel, J.J. 2009. Koa (Acacia koa) ecology and silviculture. Gen. Tech. Rep. PSW-GTR-211. Albany, CA: 

U.S. Department of Agriculture, Forest Service, Pacific Southwest 

Research Station. 129 p.

Brown, L.; Johnson, J.W. 1996. Nitrogen and the hydrologic 

cycle. Ohio State University Extension Fact Sheet AEX-463-96. 

270 Tropical Nursery Manual
Columbus, OH: Ohio State University, Food, Agricultural and 

Biological Engineering. http://ohioline.osu.edu/aex-fact/0463.

html. (June 2012).

Brundrett, M.C. 2009. Mycorrhizal associations and other means 

of nutrition of vascular plants: understanding the global diversity 

of host plants by resolving conflicting information and developing reliable means of diagnosis. Plant and Soil. 320: 37–77.

Castellano, M.A.; Molina, R. 1990. Mycorrhizae. In: Landis, T.D.; 

Tinus, R.W.; McDonald, S.E.; Barnett, J.P.; The Container Tree 

Nursery Manual, Volume 5. Agriculture Handbook 674. Washington, DC: U.S. Department of Agriculture, Forest Service: 101-167.

Evans, J. 2002. Plantation forestry in the tropics. 2nd ed. New 

York: Oxford University Press Resources. 47 p.

Habte, M.; Miyasaka, S.C.; Matuyama, D.T. 2001. Arbuscular 

mycorrhizal fungi improve early forest tree establishment. In: 

Horst, W.J.; Schenk, M.K.; Burkert, A.; Classen, N., eds. Plant 

nutrition-food security and sustainabiity of agro-ecosystems. Dordrecht, Netherlands: Kluwer Academic Publishers. 644-645 p.

Habte, M.; Osorio, N.W. 2001. Arbuscular mycorrhizas: producing and applying arubscular mycorrhizal inoculum. Honolulu, 

HI: University of Hawai‘i, College of Tropical Agriculture and 

Human. 47 p.

Janos, D.P.; Schroeder, M.S.; Schaffer, B.; Crane, J.H. 2001. 

Inoculation with arbuscular mycorrhizal fungi enhances growth 

of Litchi chinensis Sonn. trees after propagation by air-layering. 

Plant and Soil. 233: 85–94.

Keyser, H. 2002. Personal communication. Paia, HI: University of 

Hawai‘i NifTAL Project.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1989. The 

container tree nursery manual: volume 5, the biological component: nursery pests and mycorrhizae. Agriculture Handbook 674. 

Washington, DC: U.S. Department of Agriculture, Forest Service. 

171 p.

Mikola, P. 1973. Application of mycorrhizal symbiosis in forestry 

practice. In: Marks, G.C.; Kozlowski, T.T., eds. Ectomycorrhizae: 

their ecology and physiology. New York and London: Academic 

Press: 444 p. Chapter 10.

Miyasaka, S.C.; Habte, M.; Friday, J.B.; Johnson, E.V. 2003. Manual on arbuscular mycorrhizal fungus production and inoculation techniques. Honolulu, HI: University of Hawai‘i at Mānoa, 

College of Tropical Agriculture and Human Resources. 4 p.

Nitrogen Fixing Tree Association (NFTA). 1989. Why nitrogen fixing trees? NFTA 89-03: 1-2. Morrilton, AR: Forest, Farm 

and Community Tree Network (FACT Net), Winrock International. http://www.winrock.org/fnrm/factnet/factpub/FACTSH/

WhyNFT.htm.

Read, D.J. 1996. The structure and function of the Ericoid mycorrhizal root. Annals of Botany. 77: 365–374. 

Schmidt, L. 2000. Guide to handling of tropical and subtropical 

forest seed. Humlebaek, Denmark: Danida Forest Seed Centre. 

511 p.

Schmidt, L. 2007. Tropical forest seed. Berlin, Germany: SpringerVerlag. 409 p.

Singleton, P.W.; Somasegaran, P.; Nakao, P.; Keyser, H.H.; Hoben, 

H.J.; Ferguson, P.I. 1990. Applied BNF technology: a practical 

guide for extension specialists. Module Number 3: Introduction 

to rhizobia. Honolulu, Hawai‘i: University of Hawai‘i-Mānoa, 

College of Tropical Agriculture and Human Resources. 13 p. 

http://www.ctahr.hawaii.edu/bnf/Downloads/Training/BNF%20

technology/rhizobia.PDF. (April 2012).

Smith, S.E.; Read, D.J. 1997. Mycorrhizal symbiosis. 2nd ed. San 

Diego: Academic Press. 605 p.

Wall, L. 2000. The actinorhizal symbiosis. Journal of Plant 

Growth Regulation. 19: 167–182.

Wang, B.; Y.L. Qiu. 2006. Phylogenetic distribution and evolution of mycorrhizas in land plants. Mycorrhiza. 16: 299–363. 

Wilkinson, K.M.; Elevitch, C.R. 2003. Growing koa: a Hawaiian 

legacy tree. Holualoa, HI: Permanent Agriculture Resources.

Additional Reading

Alexander, I.; Selosse, M.A. 2009. Mycorrhizas in tropical forests: 

a neglected research imperative. New Phytologist. 182: 14–16.

Brundrett, M.C. 2008. Mycorrhizal associations: the Web 

resource. http://mycorrhizas.info/info.html http://mycorrhizas.

info/info.html. (February 2013).

Dawson, J.O. 2009. Ecology of Actinorhizal plants. In: Pawlowski, 

K., ed.; Newton, W.E. series ed. Nitrogen-fixing Actinorhizal 

symbioses. Dordrecht, The Netherlands: Springer: 199–227. 

Chapter 8.

Margulis, L.; Sagan, D. 1997. Microcosmos: four billion years of 

evolution from our microbial ancestors. Berkley, CA: University 

of California Press. 301 p.

Beneficial Microorganisms 271
272
273

As any experienced grower knows only too well, nursery management is a continuous process of solving problems. One recurring problem is pests. In the past, 

nursery managers waited for an insect or disease to appear and then sprayed some 

toxic chemical to wipe out the pest or disease. This approach, however, also wipes 

out natural predators of the pest, resulting in an expensive and repeating pesticide 

cycle. Instead of a knee-jerk reaction to a specific problem, “holistic” pest management is a series of interrelating processes that are incorporated into the entire 

spectrum of nursery culture. Holism is the theory that systems, and each part of a 

system, should be viewed as a whole and not as isolated parts. “Holistic,” then, is an 

approach that looks at the big picture and considers all parts. Holistic pest management is an integrated and preventative approach that considers the overall health 

of the plant and the nursery environment to prevent problems and to manage them 

wisely if they arise. The holistic approach to nursery pest management involves a 

series of four interrelated practices, which ideally function together (Wescom 1999): 

• Problem Prevention Through Cultural Measures—through good sanitation, 

proper scheduling, management of the nursery environment, and promotion 

of plant health. 

• Problem Detection and Diagnosis—through monitoring, recordkeeping, and 

accurate problem identification.

 • Problem Management—including, if necessary, timely and appropriate pest 

suppression measures and balance of pest populations with beneficial 

organisms and pest predators.

• Ongoing Process Evaluation—to learn from experience by assessment and 

improved effectiveness of pest management approaches. 

Problem Prevention and 

Holistic Pest Management

Thomas D. Landis, Tara Luna, R. Kasten Dumroese, and Kim M. Wilkinson

14

Facing Page: Beneficial insects are an important part of holistic pest management. The ladybugs (Hippodamia species

[Coleoptera: Coccinellidae]) in this photo can be seen feeding upon the psyllid pest Heteropsylla cubana (Hemiptera: Psyllidae). 

Photo by J.B. Friday.
274 Tropical Nursery Manual

Figure 14.1—A collage of biotic nursery pests. Fungal pests include damping-off (A), Phytophthora root canker (B), fungal rust on 

ō‘hi‘a leaves (C), and Botrytis blight (D). Insect pests include aphids (E), fungus gnat larvae eating seeds (F) and roots (G), leaf miners 

(H), mealybugs on palm (I), and mites making leaf galls (J). Snails and slugs (K) can damage nursery plants. Plants such as weeds, liverworts, and algae (L, M) can smother small plants. Photos A through J by Thomas D. Landis, photo K by Brian F. Daley, photo L by R. 

Kasten Dumroese, and photo M by William Pink.

B E

H

K L M

C

F I

A D

G

J
Nursery Diseases and Pests

Nurseries have many potential pests, including fungi 

(figures 14.1A–14.1D), insects (figures 14.1E–14.1J), 

nematodes, snails, slugs (figure 14.1K) and even larger 

animals such as mice and deer. Other plant species, such 

as weeds and cryptogams (moss, algae, or liverworts) 

(figures 14.1L, 14.1M) can become pests when they compete with crop plants for growing space and light.

Plant disease can also be caused by abiotic (environmental) stresses, including heat (figures 14.2A, 14.2B), 

light (figure 14.2C), and too much or too little water (figures 14.2D, 14.2E). Sometimes, people are pests when they 

misuse chemicals such as applying too much lime (figure 

14.2F) or fertilizer (figure 14.2G).

A useful concept to explain nursery pest problems 

is the “disease triangle,” which illustrates the interrelationships among the pest, host, and environment (figure 

14.3). All three factors are necessary to cause biotic disease. For example, a fungus or insect pest is able to survive inside the warm greenhouse environment and attack 

the host plant. Although many diseases may appear to 

involve only the host plant and the biological pest, environmental factors are always involved. Environmental 

stress may weaken the plant and predispose it to attack 

by the pest, or a particular environment may favor pest 

populations, enabling them to increase to harmful levels.

Abiotic disease can be visualized as a two-way relationship 

between the host plant and adverse environmental stress 

(figure 14.3). Abiotic diseases may develop suddenly as the 

result of a single injurious climatic incident, such as a heat 

wave or dust storm, or more gradually as a difficult-to-detect 

growth loss resulting from below-optimum environmental 

factors, such as a mineral nutrient deficiency.

Problem Prevention and Holistic Pest Management 275

Figure 14.2—Abiotic disease can be caused by any excessive environmental stress, 

including heat injury to germinants (A), or sunscald of mature foliage (B). Exposing 

shade-loving plants, like these Norfolk Island pine, to full sunlight can damage their 

ability to photosynthesize (C). Irrigating too much can lead to high humidity and condensation (D), which predisposes plants to many bacterial and fungal pests. Too little 

irrigation causes wilting (E), which inhibits growth. Overliming of growing media (F), 

misapplication of pesticides, or improper fertilization (G) can cause chemical injuries. 

Photos by Thomas D. Landis.

B

E

C

F

A

D

G
Problem Prevention Through 

Cultural Measures

Preventing problems is better than having to deal with 

them after symptoms occur. Prevention steps include maintaining good sanitation and hygiene in the nursery, proper 

crop scheduling to avoid diseases and health problems in 

holdover stock, and promoting plant health through good 

nursery management. 

Good Sanitation

The most logical approach to disease management is to 

prevent diseases by excluding pests from the growing area. All 

diseases are much easier to prevent than to cure. Sanitation 

begins with nursery site selection and preparation. 

A source of clean water for irrigation is one of the key considerations in site selection. Some water sources harbor damping-off fungi including Pythium and Phytophthora (Jaenicke 

1999) and need to be avoided if possible. (Treating water may 

be an option; see Chapter 11, Water Quality and Irrigation.) 

A weed-free fabric groundcover or concrete floor under nursery benches can also be instrumental in preventing problems. 

In contrast, having bare soil under plants can allow soil 

pathogens to splash or blow onto nursery plants and can be 

a favorable environment for weeds. Weeds near or under 

benches may provide cover for insect pests including thrips, 

aphids, whiteflies, and weevils. Weeds can also be a source of 

pathogenic fungi, such as Fusarium (James 2012). Excluding 

seed-eating rodents and birds with screens or other barriers 

can prevent these animals from becoming problems and also 

keeps out any diseases and pests they might bring in on their 

bodies. Vegetation adjacent to nurseries should be assessed 

to see how it might positively or negatively affect nursery 

plant health. For example, a nearby hedge consisting only of 

mahogany trees infested with the local mahogany shoot borer 

may increase the likelihood of these pests entering the nursery and affecting a crop of mahogany seedlings. Well-planned 

adjacent vegetation in hedgerows and windbreaks can provide refuge for beneficial organisms, however, including 

pollinators and pest predators such as ladybugs, predatory 

wasps, spiders, amphibians, and birds. 

Pests generally enter the nursery growing area through 

the following modes:

• Wind—Airborne fungal spores, seeds, or insects can 

be introduced on the wind.

• Water—Fungus and cryptogam spores and weed 

seeds can be introduced through irrigation water.

• Growing Media—Most growers try to use growing media components that are free from pathogens. 

However, potentially harmful fungi can sometimes be 

isolated from some growing media components. When 

topsoil is used as the growing medium, or as part of a 

mix, it can carry a variety of insect, fungal, and weed 

pests to the nursery. See Chapter 6, Growing Media, for 

more information about the risks associated with using 

soil as the growing medium.

• Containers—Reusable containers may contain 

residual growing medium or plant roots that harbor 

potentially pathogenic fungal propagules, moss, or 

algae.

• Surfaces in the Growing Area—Floors, benches, and 

other surfaces in the growing area may harbor pests.

• Propagation Materials—Seeds, transplants, or cuttings are sometimes contaminated before they reach 

the nursery. Importing plants from other nurseries 

poses a very high risk of introducing new pests. If the 

plants have been imported from other islands, this 

poses a risk not only to the nursery but to the entire 

island ecosystem as well. Imported nursery stock has 

been the culprit for introducing several invasive pests 

and plants into island ecosystems.

• Transported Pests—Infested soil or other materials 

can be carried into the growing area on tools, equipment, or the shoes of workers or visitors.

• Mobile Pests—Insects, birds, and rodents can enter 

the growing area directly.

276 Tropical Nursery Manual

Figure 14.3—The “disease triangle” illustrates the concept that 

a host, a pest, and a conducive environment are necessary to cause 

biotic disease. Abiotic disease occurs when environmental factors, 

like excessive moisture or heat, injure the host plant. Illustration 

adapted from Dumroese and others (2008).
Checklist for Preventing Diseases and Pests With 

Good Sanitation

• Start with clean seeds. Seeds can be rinsed with a 

diluted bleach or hydrogen peroxide solution before 

stratification or sowing to help prevent seed and seedling diseases (see Chapter 9, Seed Germination and 

Sowing Options).

• Remove all plant debris from the nursery area before 

sowing each crop. Also, clean tables, aisles, sidewalls, 

and floors with a mild bleach or soap solution before 

sowing.

• Regularly and vigilantly remove all weeds growing 

under benches and within the crop.

• Use containers that have been cleaned (see Chapter 7, 

Containers).

• Heat-pasteurize (steam or solarization) any growing 

media ingredients that may contain pests or diseases. 

Use soil-free media if possible (see Chapter 6, Growing Media).

• Prevent algae from forming on the floors and benches 

by ensuring rapid drainage of excess irrigation water 

and by properly managing irrigation frequency (see 

Chapter 11, Water Quality and Irrigation). Algae and 

pools of water provide a breeding ground for fungus 

gnats and shore flies and the slick surfaces can be a 

safety hazard to nursery staff.

• Use hooks to keep the hose nozzles off the floor and 

disinfect planting tools after each use.

• Keep plants off the floor and out of contact with soil 

if feasible. Elevating plants on benches, with a fabric 

groundcover or concrete floor underneath, helps to 

reduce pest problems.

• Test nursery water for problems and treat if necessary.

• Use precaution if importing plant material from other 

nurseries, especially from other islands. Imported 

plant materials should be certified pest free. You 

will ideally use only local, native, or traditional plant 

materials in your nursery to minimize the risk of 

spreading insects, diseases, and invasive plants. 

Proper Scheduling

Crop scheduling is an important component of holistic 

pest management. A typical tropical plant nursery will be 

growing a wide variety of plant sizes and species with different growth rates. For example, in Hawai‘i, koa can be 

grown from seeds to shippable size in as little as 3.5 months, 

whereas native Hawaiian sandalwood seedlings take 1 year 

or longer to be large enough to ship. Slow-growing plant 

species should be started first so they have maximum time 

to grow. Species that grow very quickly should be scheduled 

later in the season so that they do not become “top heavy” 

(figure 14.4A) or rootbound (figure 14.4B). Plants that have 

grown too large for their containers are easily stressed and 

Problem Prevention and Holistic Pest Management 277

Figure 14.4—Keeping nursery stock at the ideal size for their containers is a challenge with rapidly growing species, such as this Acacia koa, 

which quickly become “top heavy” (A). Although it is harder to detect, roots may grow so fast that they become “rootbound” (B) and 

stressed, which leaves them susceptible to fungal root diseases. Tropical plants cannot be held over for too long without becoming 

unbalanced, so crop scheduling is critical (C). Photos by Thomas D. Landis.

A B C
can harbor insects and other pests that can infect the rest 

of the crop. One of the most common problems in tropical 

nurseries is that most plants never stop growing and so it is 

challenging to keep crops from becoming too large for their 

containers (figure 14.4C). See Chapter 4, Crop Planning: 

Propagation Protocols, Schedules, and Records, for more 

information on crop scheduling.

If you are growing a new species and are unsure of the 

growth rate, contact other nurseries, and check the propagation protocols in the Native Plant Network (http://

www.nativeplantnetwork.org).

Managing the Nursery Environment

An important aspect of preventing disease is to maintain 

environmental conditions (such as moisture, light, and ventilation) that are conducive for plant growth but that do not 

favor pests or have damaging environmental extremes. For 

example, root diseases are common in all types of nurseries worldwide, and are often brought on by environmental 

stresses. Swedish researchers studied the fungal pathogen 

Cylindrocarpon destructans in relation to root rot problems 

of pine seedlings in container nurseries. Suspected predisposing stress factors included excessive moisture, low light, 

and exposure to fungicides. They found that C. destructans

typically invades dead or dying roots and then uses these 

sites as a base for invasion into healthy roots (Unestam and 

others 1989). Opportunistic pathogens, such as Fusarium, 

are more likely to cause disease when plants are under environmental stress (figure 14.5).

Marginal environmental conditions often favor insect 

pests. For example, fungus gnats become a problem only 

under wet conditions and especially in locations where algae, 

moss, and liverworts have been allowed to develop. Often, 

these conditions exist under nursery benches, where water 

can puddle and excess fertilizer promotes their growth. This 

problem is particularly common where nursery floors are 

covered by a weed barrier or gravel. Switching to concrete 

floors can often cure a fungus gnat problem because concrete dries faster and is easier to keep clean.

Checklist for Preventing Diseases and Pests by 

Managing the Nursery Environment

• During sowing and the establishment phase of the crop, 

keep humidity levels and condensation problems low 

by venting the nursery area well. Do not overwater 

germinating seeds and seedlings. During the active 

growth stage, reduce humidity within the leaf canopy 

to prevent the development of many foliar diseases. 

Reducing humidity can be accomplished by improving 

air circulation by increasing distance between plants, 

increasing the frequency of ventilation, and pruning 

shoots as necessary.

278 Tropical Nursery Manual

Figure 14.5—Many nursery diseases are caused by stresses, which predispose plants to attack by opportunistic pests. Illustration 

adapted from Dumroese and others (2008).

Water Can Be Harmful to Plant Health

Too much water on plants (figure 14.2D) encourages 

damping-off, root disease, fungus gnats, moss and liverwort growth, excessive leaching of mineral nutrients, 

potential groundwater contamination, and foliar diseases 

such as Botrytis blight (figure 14.1D), powdery mildew, 

and Rhizoctonia. Over-irrigation may cause plants to 

grow rapidly and spindly making them more susceptible 

to environmental stresses such as excessive heat, wind, or 

cold. Too little water on plants may cause damage to root 

systems through salt accumulation or desiccation, allowing entry points for root disease pests. Plants under severe 

moisture stress have lower resistance to pests and stresses 

associated with heat, humidity, and wind.

Proper irrigation management is one of the best ways 

to limit pest problems.
• Water only in the morning, never later in the day. 

Favorable environmental conditions for several fungal 

diseases include a film of moisture for 8 to 12 hours, 

high relative humidity, and warm temperatures. By 

watering early, rising daytime temperatures will cause 

water to evaporate from the leaf surfaces and reduce 

favorable conditions. (As an exception, Botrytis 

problems often occur when conditions are humid but 

temperatures are cool [James 2012].)

• To optimize healthy environmental conditions for 

different crops, use separate propagation structures 

for growing plants with different environmental and 

cultural requirements, or, if you have a single growing 

area, group plants with similar growing requirements 

together and take advantage of microenvironments 

within the nursery area. For example, north of the 

equator, the south side of the nursery is usually 

warmer and drier than the north or east section of 

the nursery. Plants requiring drier conditions will 

benefit from being grouped on the south and west 

sides of the nursery. Plants requiring cooler temperatures (such as high-elevation species) or those 

requiring more frequent irrigation can be grouped 

together on the north and east sides.

• Improve air circulation and reduce humidity and 

condensation with spacing between plants, using 

fans, and orienting to prevailing breezes to produce 

horizontal airflow. 

Promotion of Plant Health

Healthy plants are more able to resist infection from 

pathogenic fungi and attack from insects and other pests 

and can also tolerate environmental stresses better. Much 

of this resistance can be attributed to physical characteristics such as a thick, waxy cuticle on the foliage, balanced 

shoot and root, and optimum mineral nutrient contents. 

Some healthy plants also produce chemicals such as 

tannins, which deter pests. Management of the nursery 

environment to ensure plants receive proper sunlight 

and water as described previously, as well as the steps of 

good sanitation and proper scheduling, is important for 

plant health. Appropriate nutrition, growing medium, 

and container size and type; use of beneficial microorganisms and other practices described in this manual are 

important for promoting plant health on all levels.

Plants must have a proper growing medium to be 

healthy. Good growing media provides physical support 

and aeration to the roots, and holds water and nutrients 

for the plant, as described in Chapter 6, Growing Media. 

Appropriate container selection is described in Chapter 

7, Containers. Different species will require different 

containers types and sizes based on their growth characteristics and target specifications. Fertilization at the 

appropriate level to help plants grow in a healthy and 

balanced way is discussed in Chapter 12, Plant Nutrition 

and Fertilization. Excessive fertilization can make plants 

more susceptible to problems, including sucking insects 

and diseases. Consider eliminating nitrogen fertilization 

during germination because it encourages damping-off 

fungi (James 2012).

If plants do become sick or unhealthy, it is rarely, if 

ever, practical to try and “save” them. Remove dead and 

dying plants; dispose of them away from the nursery to 

prevent reinfection. Remove any plant debris in the containers or on the floor on a regular basis.

Problem Detection and Diagnosis

Even with good sanitation, crop planning, plant health, 

and management of the nursery environment, problems 

may appear. A holistic approach emphasizes detecting problems early through regular monitoring and recordkeeping 

so they can be addressed quickly and with minimal cost and 

effort. If a problem is detected, the time is taken to diagnose 

it accurately and determine its true source. With an accurate 

diagnosis, the problem can then be treated accordingly. 

Monitoring and Recordkeeping

Regular monitoring or “scouting” is a critical part of 

the holistic approach. A daily walk-through of the nursery will reveal developing pest outbreaks or environmental problems that are conducive to pests while problems 

are still minor and easily corrected. In small nurseries, 

the grower or nursery manager should do the inspections, but, in larger facilities, one person should be designated as the pest scout. This person should have nursery 

experience and be familiar with all the plant species at 

all growth stages. It is essential to know what a healthy 

plant looks like before you can notice any problem. It is 

also essential to know what beneficial organisms look 

like (such as natural enemies of pests, mycorrhizal fungi 

growing on the roots, and so on). The person should also 

be inquisitive and have good observation skills. Often, 

irrigators serve as pest scouts because they are regularly 

in the nursery, checking whether plants need watering.

Crop monitors (figures 14.6A, 14.6B) need to carefully 

inspect each species being grown, record the environmental conditions in the growing area, and make other 

observations. It is important to establish a monitoring 

and recordkeeping system for all areas of the nursery, 

including any structures, rooting chambers, and outdoor 

Problem Prevention and Holistic Pest Management 279
growing areas. These records are invaluable for avoiding 

future pest problems when planning the next crop. In 

addition to looking at crop plants, pest scouts should check 

all nursery equipment and should carry a 10X to 20X hand 

lens for close examination and a notebook or voice recorder 

to record observations. When a problem is noted, a camera 

with a closeup lens is an excellent way to document the problem (figure 14.6C). All observations should be recorded in a 

daily log (figure 14.6D), and any suspicious problems need 

to be immediately reported to the nursery manager. Inspections need to occur on a daily basis after sowing the crop and 

during the establishment phase, when plants are most susceptible to diseases such as damping-off. When problems are 

detected early, plants can be treated or isolated from the rest 

of the nursery or greenhouse crop.

Use yellow sticky cards (figure 14.7) to detect white flies, 

aphids, fungus gnats, and shore flies. Place one to four cards 

every 1,000 ft2 (90 m2) and space them evenly in a grid pattern, with extra cards placed near vents and doorways. 

Inspect these cards weekly to detect and monitor these pests. 

Record the information and replace the cards as needed. Use 

blue sticky cards, which are more attractive to thrips, around 

plants susceptible to this pest. Observing and recording pest 

predators can also be valuable: insect-eating birds, amphibians, spiders, predatory wasps, and other creatures that may 

be residing in or near the nursery.

Accurate Problem Identification

Nursery managers and pest scouts must be able to 

identify problems quickly and accurately before the 

problems can cause significant damage. Although biological pests such as fungi and insects are always present, 

abiotic stresses typically cause more problems. Disease 

diagnosis requires a certain degree of experience and training, and nursery workers need to be trained to quickly spot 

280 Tropical Nursery Manual

Figure 14.6—Regular pest monitoring or “scouting” is critical to identifying problems early (A). Pest scouts inspect the seedlings and 

the nursery environment (B). Pest scouts should carry a hand lens, notebook or recording device, and camera to identify and document 

problems (C). All potential problems need to be noted in the daily log (D) and reported to the nursery manager. Photo A by Tara Luna, 

photo B by Diane L. Haase, photo C by Thomas D. Landis, and photo D by Kim M. Wilkinson.

Figure 14.7—Yellow sticky cards are essential for monitoring 

the types and population levels of insects pests. Photo by Thomas 

D. Landis.

A B C

D
new problems and incidents of abiotic injury. Workers who 

are in the growing area daily have the best chance of detecting potential problems so they can be accurately identified 

and treated before they can intensify or spread.

A general characteristic of abiotic diseases is that the 

symptoms tend to show up very quickly (see textbox example). Problems caused by biotic agents generally take longer 

to develop, with decline occurring over time (James 2012). If 

abiotic damage has been eliminated as the cause of the problem, then a pest or disease must be diagnosed. Landis and 

others (1990) features identification keys (figure 14.8), and 

color photographs of many diseases, insect pests, and horticultural problems of conifer crops, but no single reference 

is available about diseases and pests of tropical plant species. 

Although pest scouts can make a tentative diagnosis of disease and pest problems, they need to confirm their conclusions with the nursery manager and a trained nursery pest 

specialist. Plants with undiagnosed symptoms may be sent 

whole to the nearby extension office or land-grant university 

for diagnosis and suggested followup.

Many diseases and insect pests damage a wide variety 

of host plants. For example, damping-off fungi (figure 

14.1A) affect all species during germination and emergence 

and Phytophthora root rot (figure 14.1B) can infect larger 

plants. Other pests are host specific, however, and pest 

scouts must understand their basic biology and life cycle 

for accurate diagnosis. In tropical nurseries, it is very important for nursery growers to be aware of newly introduced diseases and the range of species that are affected by them. For 

example, ‘ōhi‘a rust is caused by a fungus (Puccinia psidii), 

which also infects guava, eucalyptus, and a range of native 

species in the Myrtle family. In Hawai‘i, it has been found 

on introduced timber and ornamental species and native and 

endangered species of nīoi. It affects leaves and meristems, 

inhibiting growth and development and is especially severe 

on seedlings, cuttings, and saplings. The first symptoms are 

chlorotic specks, which after a few days become pustules 

containing uredia producing yellow masses of spores (figure 

14.1C). Pustules can coalesce and parts of the plant can be 

completely covered with them. After about 2 to 3 weeks, pustules dry and become necrotic. The disease can cause deformation of leaves, heavy defoliation, dieback, stunted growth 

and eventual death. On fruits of guava, and other hosts in the 

Myrtle family, the lesions occur mostly on buds and young 

fruits that eventually rot as the rust matures. Understanding the life cycle of the pest is essential for accurate diagnosis 

and management. Some of the more common tropical plant 

pests and ways to monitor, diagnose, prevent, and, if necessary, treat them, are included in table 14.1.

Problem Prevention and Holistic Pest Management 281

Abiotic Factors as “The Usual Suspects”

I was growing a new crop of a mahogany species. The plants were fairly young with only four to six leaves each. 

They were growing slower than past crops, and some of the oldest leaves had brown leaf margins that I attributed 

to uneven watering. Otherwise the seedlings seemed fine. Because they were growing slowly, I thought I would 

help them along by applying a fertilizer. But a day or two after that, the seedlings were all wilting and the brown 

leaf margins were spreading into the leaves. With the leaf margin burning and the plants wilting and dying, I suspected root rot. I immediately sent some affected plants to the local land grant university extension service, with 

an urgent message. While I waited for the reply, I researched options for treating root rot, most of which involved 

safety gear and unattractive pesticides. The extension agent contacted me sooner than I had hoped, saying he had 

not completed his diagnosis but so far had detected dangerously high levels of salts in the growing medium; the 

plants were exhibiting signs of salt toxicity. As we talked, I remembered that I had used a batch of untested coir as 

one of my growing medium components for this crop. I had heard that coir can sometimes contain high levels of 

salts because of processing, but I had not taken the time to test this batch before planting. The seedlings had been 

struggling along in somewhat salty conditions (which explained the slower growth and burned leaf margins) and 

my fertilizer application added even more salts, pushing them over the edge into salt toxicity. With that, they could 

not take in enough water with their roots because of salts in the medium, and the chloride they were taking up was 

scorching their leaves. The extension agent suggested I flush out the salts from the growing medium. As soon as I 

got off the phone, I drenched the seedlings with clean water until water dripped out the bottom of the containers, 

and repeated again a few hours later. Almost overnight, the seedlings perked up. Some of the smallest had died and 

were a loss, but the remaining trees quickly grew new leaves and within a few weeks were in full health. I learned 

from this scare that abiotic factors need to be the first suspects when diagnosing plant problems. A quick check 

from an EC meter would have revealed the problem inexpensively in only a few minutes. I also learned that taking 

the time to ensure the basics, such as growing medium quality, are okay can save a lot of stress—for me and for the 

plants. — Kim Wilkinson
Check with local regulations before ordering and introducing biological control organisms.

Problem Management

After a problem is accurately diagnosed, it must be 

evaluated to determine what, if any, management action is 

necessary. The first line of defense is always to assess and, if 

possible, improve pest prevention practices including good 

sanitation, physically excluding pests, proper crop scheduling, and promotion of optimal plant health through 

appropriate fertilization, irrigation, ventilation, and 

other care. If the problem persists, the holistic approach 

calls for assessing the effects of the pest or disease. In some 

cases, simply doing nothing may be the most cost-effective 

solution; plants can tolerate some damage and nature can be 

allowed to take its course (Mizell and Short 2009).

If action is taken, the goal is usually to reduce crop 

losses to acceptable levels, rather than to eradicate pests. A 

spectrum of management options is available, with treatments including—

• Modifying and managing the nursery environment.

• Culling affected seedlings.

• Mechanically removing pests (picking off, swatting, 

trapping—ensure staff are trained to know the 

difference between beneficial pest predators and 

actual pests).

• Using biological controls such as beneficial microorganisms and pest predators, diseases, or parasites 

(table 14.1).

• Using the least toxic pesticides available as a last 

resort to lower pest populations to a tolerable level 

(Wescom 1999).

Timely and Appropriate Pest 

Suppression Measures

Synthetic or natural pesticides should be considered 

only as a last resort after other environmental and horticultural control measures have been considered. Practicing 

good hygiene and sanitation, including using pathogenfree seeds, growing media, and containers, is critical. You 

can often reduce or even eliminate the use of pesticides 

by physically excluding pests with screens or barriers that 

can keep out pests large and small, from insects to rats to 

pigs (figure 14.9). If seedlings are affected by fungi, you 

can reduce the seedling’s susceptibility by spacing out the 

plants and increasing air movement through the nursery area, and by watering appropriately and lowering the 

humidity around the plants (Wescom 1999).

282 Tropical Nursery Manual

Figure 14.8—Pest identification keys are very useful in helping make the correct diagnosis of nursery pests. Illustration from Dumroese 

and others (2008).
Problem Prevention and Holistic Pest Management 283

Table 14.1—Common pests of tropical plants and their symptoms, prevention, and treatment. Although a variety of pesticides are 

available for these diseases, only biological and organic control options are provided. Ideally, nursery staff will monitor daily for signs 

of these diseases.

Type of pest Signs and symptoms Monitoring Prevention Biological 

control options

Fungi

Botrytis blight 

(Botrytis cinerea)

Plants have leaf blights, stem 

cankers, gray mold. Mold is 

most noticeable on older foliage. 

Affected seedlings are usually 

in groups.

Concentrate monitoring 

where crop is closely spaced 

and air circulation is poor 

and at the base of plants. 

Look for dieback, stem 

cankers, and powdery gray 

mold on foliage.

Increase spacing between 

containers as crops grows larger. 

Water only in the morning or use 

subirrigation. Keep humidity low 

by increasing air flow. Surface 

sterilize greenhouse floors, walls, 

and benches between seedling 

crops.

Apply products containing 

Trichoderma harzianum or 

Streptomyces griseoviridis to foliage. When discovered, remove 

symptomatic seedlings from 

growing area.

Fusarium wilt 

(Example: 

Fusarium oxysporum)

Leaves cup downward or stems 

bend in a crook. In later stages, 

brown or red streaks can be seen 

on the leaves. Orange spores 

(sporodochia) may be on stem.

Look for downward bending 

leaves or “cupping” of leaf 

margins. Can be confused 

with water stress or root rot. 

Send sample to laboratory to 

confirm.

Use mesh benches to encourage 

airflow. Do not overwater crop. 

Keep humidity low. Sterilize reused containers between seedling 

crops. Use pathogen-free seeds 

and growing media.

Apply products containing 

Trichoderma harzianum or 

Streptomyces griseoviridis as a 

soil drench. Remove and isolate 

infected plants as soon as problem is detected.

Crown rots 

(Example: 

Fusarium species)

Plants are stunted, wilted, and 

off-color. Roots are discolored 

and turn brown or black. Main 

stem becomes weak and water 

soaked in appearance. Some 

affected seedlings may have 

orange-yellow spore structures 

[sporodochia] on main stem just 

above the ground line.

Monitor weekly for wilted, 

off-colored plants with 

discolored root systems. 

Pay attention to media that 

stays wet. Affected seedlings 

are usually concentrated 

in groups. Send samples 

to diagnostic laboratory to 

confirm pathogen identity. 

Do not overwater crop. Increase 

spacing between containers as 

crop grows larger. Keep humidity 

low.

Apply products containing 

Trichoderma harzianum, 

Trichoderma virens, or 

Streptomyces griseoviridis as 

a drench.When discovered, 

remove symptomatic seedlings 

from growing area.

Root rots 

(Examples: species of 

Fusarium, Pythium, 

Rhizoctonia, 

Phytophthora, and 

Cylindrocarpon)

Seedlings are stunted, wilted, 

chlorotic, or necrotic and 

often occur in groups. Roots 

are discolored (dark brown or 

black) and decayed (epidermal 

tissues may be easily sloughed 

off). Main stems become weakened and water soaked. Some 

affected seedlings may have 

orange-yellow spore structures 

[sporodochia] on the main stem 

just above the ground line.

Monitor weekly for wilted, 

chlorotic, and necrotic 

seedlings with discolored 

and decayed roots. Problems 

can be especially severe in 

containers with prolonged 

wet growing media (poorly 

drained). Affected seedlings 

are usually concentrated 

in groups. Send samples 

to diagnostic laboratory to 

confirm pathogen identity. 

Do not overwater crop. Increase 

spacing between containers as the 

crop grows larger. Keep humidity 

low. Sterilize re-used containers between seedling crops. Use 

pathogen-free seeds and growing 

media.

Apply Trichoderma harzianum, 

Trichoderma virens, or 

Streptomyces griseoviridis as 

a drench.

Damping-off

(Examples: species of

Fusarium and 

Pythium)

Seeds do not germinate or 

seedlings collapse at soil line 

just after emergence. Dark 

decayed spots appear on stems 

at soil line of emerged seedlings. 

Surviving plants may later 

develop crown or root rot.

Monitor new sowing daily 

during germination and 

establishment phases. 

Excavate seeds and cut to 

check for decay and discard 

infected containers immediately.

Cleanse seeds and growing area. 

Use sterile media and containers. 

Avoid over-sowing, crowding of 

seedlings, or planting seeds too 

deeply. Keep greenhouse and 

media temperatures warm during 

germination and establishment. 

Keep humidity low. Do not fertilize with nitrogen during seedling 

establishment phase.

Treat seed with running-water 

rinses for a minimum of 48 

hours prior to sowing, or surface 

sterilize with bleach or hydrogen 

peroxide solutions (1:10). Apply 

products containing Trichoderma harzianum or Trichoderma virens as a drench at time 

of sowing.

Powdery mildew 

(Examples: species of

Blumeria, 

Podosphaera, 

Microsphaera, and 

Erysiphe)

Plants may have white powdery 

fungal growth on upper or lower 

leaf surfaces. If severe, a white 

coating can be seen on foliage.

Monitor weekly. Inspect 

susceptible species. Look 

in areas near vents and 

greenhouse entrances or any 

location with a sharp change 

between day and night temperatures. Use a hand lens to 

see white, powdery threads 

and spores.

Place susceptible species where 

drastic changes in temperatures 

do not occur. Water only in the 

morning or use subirrigation. 

Keep humidity low. Increase spacing between containers as crop 

grows larger.

Remove infected leaves as soon 

as detected. Move infected plants 

to structure with more constant 

temperatures. Treat with Neem 

oil or horticultural oil. Try test 

tray first. Can also use sulfur 

powder as an organic preventative fungicide. Some plants are 

sensitive to sulfur injury so use 

lowest rate recommended. Do 

not apply within 2 weeks of an oil 

spray treatment.
284 Tropical Nursery Manual

Type of pest Signs and symptoms Monitoring Prevention Biological 

control options

Rhizoctonia web blight 

(Example: 

Rhizoctonia solani)

Stems and leaves may collapse 

and turn to mush with fine, 

web-like fungal strands on the 

plant tissue and at soil line.

Monitor leafy herbaceous 

plants, especially where they 

are closely spaced. Look for 

cobweb-like growth that 

mats leaves. Send samples 

to diagnostic laboratory to 

confirm pathogen identity.

Sterilize containers between 

seedling crops. Use mesh 

benches to encourage airflow. 

Place susceptible crops near 

vents and fans. Increase spacing 

between containers as crop 

grows larger. Keep humidity low.

Apply products containing 

Trichoderma harzianum as a 

preventative.

Fungal rusts

(Examples: species of

Gymnosporangium, 

Cronartium, 

Peridermium, and 

Melampsora)

Rust brown spots or stripes may 

be seen on lower and upper leaf 

surface.

Monitor regularly and be 

sure to check undersides of 

foliage. 

Group susceptible species where 

temperature and humidity can

be easily controlled. Increase 

spacing between containers as the

crop grows larger. Keep humidity 

low. May have to eliminate 

rust-alternate hosts if present in 

the vicinity of the nursery.

Isolate plants immediately.

Fungal leaf spots

(Examples: species of

Alternaria, Septoria, 

Botryosphaeria, 

Taphrina, Rhytisma, 

and Phyllosticta)

Alternaria leaf spots are usually 

brown or black with a yellow 

border. Septoria leaf spots are 

small gray to brown with a dark 

brown edge.

Monitor weekly for leaf 

spots. With a hand lens, look 

for small fungal fruiting 

bodies. Send samples to 

diagnostic laboratory to 

confirm pathogen identity.

Use mesh benches to encourage airflow. Keep nursery floor 

clean and free of pooled water. 

Water only in the morning or use 

subirrigation. Do not overwater 

crop. Keep humidity low. Increase 

spacing between containers as 

crop grows larger.

Periodically remove affected 

seedlings from growing areas. 

Apply products containing 

Trichoderma harzianum as a 

preventative. Remove infected 

leaves as soon as problem is 

detected. Isolate infected trays of 

plants from the rest of the crop.

Insects

Sucking insects

(Example: Aphids)

Plants have distorted new 

growth, sticky honeydew, and/or 

sooty mold.

Monitor twice weekly. Look 

on underside of leaves and 

on tips of new stems.

Don’t overfertilize. Shoot prune 

vigorous tender growth as 

needed. Watch for outbreaks.

Use aphid midges, aphid 

parasites, or lady bugs. Apply 

insecticidal soap every 6 days. 

Pyrethrin-based products can 

also be effective as a contact 

insecticide.

Fungus gnats 

(Example: 

Bradysia species)

Seeds that do not germinate 

or plants with weak or stunted 

growth and root damage.

Monitor every other day, 

especially during germination and establishment 

phases. Look for tiny winged 

flies near growing media 

surface. Use yellow sticky 

cards to detect adults.

Keep nursery floor clean and free 

of pooled water and algae. Do not 

overwater crop. Use a good seed 

mulch.

Use yellow sticky cards to reduce 

adult population levels. Apply 

products containing Bacillus 

thuringiensis ssp. israeliensis 

every 7 days as a drench. Formulations of entomopathogenic 

or insecticidal nematodes in the 

genera Steinemema and Heterorhabditis are also effective.

Mealybugs Plants may have white cottony 

residue. Sticky honeydew on 

leaves and sooty mold may 

develop.

Look for small, oval, softbodied insects covered with 

a white, wax-like layer on the 

underside of leaves.

Keep nursery environment 

clean. Increase spacing among 

plants. Practice good sanitation. Keep temperatures cool. 

Remove infected plants.

Apply Cryptolaemus montrouzieri. Use predatory beetles or 

parasitic wasps; pyrethrin insecticides are effective but must 

contact the pests. Spray with 

insecticidal soap or oil, or swab 

bugs with rubbing alcohol (limit 

alcohol contact with foliage).

Soft scales Honeydew and sooty mold 

develop if scales are present.

Look for yellow brown to 

dark brown scale insects 

along veins and stems.

Provide plants with good growing 

conditions and proper care, especially irrigation, so they are more 

resistant to scale damage.

Use parasitic wasps, insecticidal 

soap, or pyrethrins. Prune 

infested branches or cull entire 

plant.

Spider mites Plants may have light-yellow 

flecking of leaves, discolored 

foliage. Leaf drop and webbing 

occur during outbreaks and 

severe infestation.

Look on undersides of 

leaves, especially along veins. 

Use a hand lens to look for 

webbing, egg clusters, and 

red adult mites. Look in 

areas of that are hot and dry, 

near the heaters and vents.

Lower nursery temperatures and 

raise humidity levels, especially 

in the south and west edges of 

the nursery and near vents and 

furnaces.

Use predatory mites or predatory 

midges. Apply insecticidal soap 

every 6 days.

Table 14.1—Continued
Problem Prevention and Holistic Pest Management 285

Type of pest Signs and symptoms Monitoring Prevention Biological 

control options

Thrips Plants may have distortion of 

new leaves, buds, and shoot tips. 

White scars on expanded leaves.

Use blue or yellow sticky 

cards placed just above 

canopy foliage for detection.

Increase container spacing on 

leafy crops as needed to detect 

problems early.

Use predatory mites, pirate bugs, 

lacewings, insecticidal soap, and 

pyrethrins.

White flies Plants may have distorted new 

shoot and leaf growth.

Use yellow sticky cards to 

detect adults. Look for adults 

on the uppermost tender 

leaves. Immature larvae are 

found on the underside of 

leaves.

Inspect new plants coming 

into the greenhouse and reject 

infested plants. Create a “hostfree” period in the growing area 

for 2 weeks to starve the flies. 

Practice good weed control in and 

around the nursery.

Use predatory beetles or whitefly 

parasites. Apply insecticidal soap 

every 7 days. Pyrethrin-based 

products can also be effective as 

a contact insecticide.

Stem or twig borers

(Example: species of

mahogany shoot 

borer, twig borers [Xylosandrus compactus and 

Xyloborus species])

Affected seedlings usually have 

main stem cankers that may 

cause stem breakage at point of 

insect activity. Cankered tissues 

are usually black and insect frass 

may be present.

Look for main stem cankers 

often associated with 

blackened tissues and frass 

from insect activity. Damage 

can be randomly spread 

throughout seedling growing 

areas.

Don't allow infested stands of tree 

hosts near nursery area. Avoid 

holding susceptible seedlings past 

readiness date. 

Remove damaged seedlings 

when they are found.

Caterpillars Foliage with chewing damage; 

moths may be visible around 

plants. 

Inspect plants regularly.

Look for fecal droppings, 

bites taken out of leaves, and 

webbing (tent caterpillars). 

Check in evening or at night; 

some species hide during 

day.

Install screens around greenhouse 

to keep moths out.

Apply products with Bacillus 

thuringiensis ssp. kurstaki, 

as needed. Pyrethrin-based 

products can also be effective as 

a contact insecticide.

Other biotic pests

Bacterial diseases 

(Examples: species of 

Pseudomonas, 

Xanthomonas, and

Agrobacterium)

Plants are stunted with swollen 

or misshapen leaves. Look 

for water-soaked leaf spots or 

angular lesions on the stems. 

May have gall-like growths on 

main stem at or just below the 

ground line.

Inspect new plants coming 

into nursery. Look for 

water-soaked, dark brown to 

black leaf spots on leaves and 

wilted stem tips. Confirm 

diagnosis with a laboratory.

Increase spacing between 

containers as crop grows larger. 

Water only in the morning or use 

subirrigation. Keep humidity low.

Remove infected leaves as soon 

as problem is detected. Isolate 

infected trays of plants from the 

rest of the crop.

Nematodes 

(Examples: species of

Xiphinema, 

Meloidogyne, 

Pratylenchus, and 

Radopholus similes)

Affected seedlings will be 

stunted, with chlorotic or 

necrotic foliage; roots will be 

damaged with evidence of feeding [lesions] or galls.

Look for chlorotic and 

necrotic seedlings, often 

concentrated within groups. 

Roots will appear damaged 

or have galls.

Use soil-free media. Keep plants 

at least 3 feet off the ground on 

benches, with a groundcover or 

concrete floor underneath. 

Prevent soil from entering 

nursery. Formulations of 

entomopathogenic or insecticidal nematodes in the genera 

Steinemema and Heterorhabditis are available as biological 

control agents.

Slugs Plants may have chewed holes 

on leaves with smooth edges 

and slime that dries into silvery 

trails on foliage.

Look for chewed holes on 

leaves and trails of slime. 

Slugs hide under dense 

foliage and under containers 

and benches.

Keep plants on raised benches 

or pallets. Space containers 

as needed so that slugs can be 

detected easily.

Pick slugs off plants. Keep 

containers on benches. Use 

saucers filled with beer to attract 

slugs away from plants. Put 

down a piece of plywood or wet 

cardboard at night; slugs will 

gather underneath. Ducks will 

eats slugs.

Viruses Look for mosaic patterns on 

foliage, leaf crinkle or distortion, 

streaking, chlorotic spots, and 

distinct yellowing of veins and 

stunted plants.

Monitor weekly. Inspect 

all incoming plants. Send 

samples to laboratory to 

confirm.

Usually not a problem with native 

plants; can be a problem on 

cultivated varieties, ornamentals, 

plants grown by tissue culture.

None. Remove and discard all 

infected plants immediately. 

Thoroughly clean area of nursery 

where infected plants were 

growing.

Table 14.1—Continued
In contrast, using a pesticide even once can put you on 

a treadmill of escalating use and collateral damage that can 

be difficult to escape. For example, if you have a rat problem, you can easily and cheaply poison the rats with a rat 

poison (rodenticide). The rats that ingest this poison will 

die over the next few days. The poison, however, may move 

through the food web to natural enemies of rats, including wild raptors and owls, and to domestic or feral cats and 

dogs. The few rats that survive the poisoning can reproduce 

much more quickly than their slower reproducing natural 

predators can. In other words, a time lag will occur between 

when the rat population recovers and when their predator 

populations recover. During this time lag, another infestation of rats can emerge, with numbers now unchecked by 

scarcer, slower recovering predator populations. A reinfestation may compel you to poison the rats again, setting 

the struggling predator population back even further, if 

not eliminating them locally. This cycle could be avoided 

by considering alternate measures, such as finding ways 

to prevent the rats from entering the nursery area in the 

first place. If full exclusion is not feasible, trapping the 

rats and killing them yourself can bring their population down to a tolerable level without poison, or you can 

adopt a skillful cat or two to do the rat-trapping and killing for you. These options are more labor-intensive in 

the short term, but can pay dividends in the long term 

by keeping natural checks and balances healthy. This 

example uses large animals such as rats, owls, and cats to 

illustrate a concept, but the same population dynamics 

apply with many insects. For example, aphids multiply 

more rapidly than the ladybug predators that can help 

keep aphid populations in check, as long as the ladybugs 

are not indirectly poisoned by the nursery manager’s 

attempts to control the aphids (Hemenway 2009). 

Most tropical plants can be grown without fungicides 

and insecticides. Because the nursery environment is so 

favorable for many pests, however, chemicals are sometimes used. If choosing to use chemicals, the important 

thing is to choose a pesticide that is least toxic, safest to use, 

targeted for the specific pest, short-lived in the environment, and will not contaminate the environment or reduce 

populations of pest predators (Mizell and Short 2009). The 

chemical must be registered for the pest and crop plant 

being treated and used as the label directs.

Many people believe that any natural or organic pesticide is always safe to use, but a number of registered botanical insecticides can be toxic to applicators or the environment. The relative toxicity rating for any chemical is known 

as the LD50, which indicates the lethal dose that is required 

to kill 50 percent of a population of test animals. As can be 

seen in table 14.2, insecticides from natural sources can be 

as hazardous as chemical pesticides.

Still, if considering pesticides, we recommend that 

you always consider natural pesticides first because they 

tend to degrade faster than synthetic pesticides, which 

reduces their effect on nontarget organisms. Most natural pesticides are more likely to meet the holistic criteria 

of lower toxicity, pest-specific, short-lived, and least collateral damage compared with synthetic pesticides. The 

drawbacks of natural pesticides are that they must be 

applied more frequently because of their rapid degradation, commercial ones are often more expensive and may 

be more difficult to obtain, homemade ones may vary in 

efficacy and safety, and little data exist on their effectiveness and long-term toxicity.

286 Tropical Nursery Manual

Figure 14.9—Large-seeded species 

can be especially vulnerable to pests 

such as mice and birds after they are 

sown in containers. Measures such as 

caging newly seeded containers and 

young seedlings (A) or adding mesh or 

screen walls (B) can be used to exclude 

a variety of pests from insects to mammals. Photo A by Tara Luna, and photo 

B by Thomas D. Landis.

A B
Balancing Pest Populations 

With Beneficial Organisms and 

Pest Predators

Holistic management involves using beneficial fungi, 

insects, and other organisms to help prevent pest effects 

on crop plants (table 14.1). When these methods are 

used, it is important to maintain a healthy biological 

balance in the nursery.

Microsymbionts

As described in Chapter 13, Beneficial Microorganisms, 

microsymbionts such as mycorrhizal fungi, rhizobia, and 

Frankia have many benefits to the host plant, including 

protection against root pathogens. For example, rhizobia 

bacteria in nitrogen-fixing plants can help reduce or eliminate nitrogen fertilizer use for those crops, reducing risks 

of attracting sucking insects with overfertilized succulent 

foliage. A study of tropical hardwood nurseries in Haiti 

reported that “trees inoculated with the proper microsymbiont are more vigorous and have far fewer diseases 

or nutrient deficiency problems” (Josiah and Allen-Reid 

1991: 56). Inoculants included manufactured rhizobia for 

nitrogen-fixing legumes, crude Frankia inoculants made 

from nodules for ironwood (Casuarina species), and locally 

gathered ectomycorrhizae for pines (Josiah and Allen-Reid 

1991).

Biological Disease Control

Biological disease control uses beneficial or benign 

organisms to help protect nursery plants from diseases. 

Biological controls must also be registered for the crop, 

pathogen, and location where they will be used. Biologicals are generally designed for a particular plant pathosystem and may not have efficacy for others, although 

these products are sometimes advertised or applied as 

broad-spectrum use (James 2012). For example, biological fungicides are made of beneficial or nonharmful 

organisms applied to the growing medium to protect 

roots from certain pathogens (Francis 2009). Different 

types of biological fungicides have different ingredients 

and modes of action, some preventing pathogen infestations in crops by competing with or excluding pathogens, and others by producing antagonist, antifungal, or 

antibiotic substances to inhibit pathogens. Some of these 

products are certified for organic production, while others are patented, hybrid organisms with specific modes 

of action. Growers in tropical areas may wish to explore 

the concept of biological disease controls and see if 

locally appropriate, naturally occurring options for disease control are feasible to make or purchase. Introduction of new microorganisms is usually not appropriate 

or legal for island ecosystems. Also, effects on pathogens 

are generally much more subtle than chemical pesticides. 

A few examples of biological fungicides are included in 

table 14.1.

Ecosystem Balance

 Only a few species of insects, bacteria, and fungi are 

harmful to plants. Most are harmless or beneficial, acting as pollinators, nutrient recyclers, food for birds and 

mammals, and pest predators (figure 14.10). The use of 

broad-spectrum chemical pesticides can be deadly to 

many harmless and beneficial organisms and to some 

of the wildlife that eat them. Furthermore, widespread 

herbicide use has enabled people to annihilate supposedly “useless” vegetation in hedges, gulches, edges, and 

fallow areas—destroying much of the food and shelter 

needed for diverse life forms. With the loss of ecological 

balance caused by these indiscriminant practices (Hemenway 2009), crop losses have been increasing. Reversing 

the loss of biodiversity and ecological balance on a local 

level is beneficial for nurseries and for the environment.

Practicing holistic pest management avoids harming 

beneficials with indiscriminant pesticide use. It is important that everyone who works in the nursery recognizes 

and protects benign and beneficial critters. Uninformed 

people may see a harmless spider and react by squishing it unless they understand how that spider might be 

helpful. Insect pest predators can include spiders, beetles, lacewings, wasps, Syrphid flies, pirate bugs, snakes, 

birds, toads, lizards, and more (Josiah and Allen-Reid 

1991). In addition, parasitic insects exist, such as parasitic wasps and tachinid flies, that kill pests by laying 

eggs inside them (Hemenway 2009). 

Problem Prevention and Holistic Pest Management 287

Insecticide Class

Toxicity 

rating 

(oral LD50 in 

mg/kg)

Label warning 

(danger is most 

toxic and caution is 

the least)

Nicotine Botanical 50 to 60 Danger

Sevin Synthetic 850 Warning or caution

Malathion Synthetic 885 to 2,800 Caution

Pyrethrin Botanical 1,200 to 1,500 Caution

Neem Botanical 13,000 Caution

Table 14.2—Comparative safety of common botanical and 

synthetic insecticides. Adapted from Cloyd (2004).

LD50 = lethal dose to 50% of subjects.
Nurseries may also protect and create habitat for beneficials and foster biodiversity, from soil organisms to 

insects, birds and, where appropriate, larger mammals. 

These “refuges” can include a simple patch of native 

flowers planted around the nursery’s entrance sign, a 

border planting at the edge of the groundcover, a fallow area, windbreaks, hedgerows, or dedicated reforestation areas. For example, windbreaks have been found 

to harbor beneficial insects and birds that help to balance out populations of crop pests, reducing needs for 

pesticides (Stace 1995). Windbreaks on farmlands have 

been shown to harbor not only native wildlife, but also to 

foster native tree seedlings and ecosystem regeneration 

processes within the windbreak (Harvey 1999). 

A key to planning refuge areas near the nursery is 

to consider what sorts of life forms you want to attract. 

Then, consider what they need for food and forage, safe 

travel corridors, reproductive habitat and nesting sites, 

and shelter from weather and predators. For example, 

native bees are important pollinators for native plants 

in many parts of the world, and their populations have 

been in decline. Different bees have different needs, but 

most native bees require a year-round supply of pollen 

and nectar from a diversity of native flowering plants. 

For shelter, native bees need a safe place to nest, usually 

in undisturbed ground, in hollow stems, or holes in trees 

(Vaughan and Black 2008). 

The tips below can help to attract a diversity of benign 

and beneficial insects, animals, and other organisms to 

available areas including borders, hedges, windbreaks, 

and other refuge spots around the nursery (adapted from 

Wilkinson and Elevitch 2000):

• Learn about the intended species’ needs for food, 

habitat, and other necessities. Plant the known food 

sources for target species. Native plant species are 

most likely to support native life, from soil fauna to 

birds. Some native wildlife may also have become 

accustomed to exotic species for food or habitat as 

well, however, and some naturalized exotic species 

may be important predators for exotic pests.

• Do not practice “clean culture.” As long as it does 

not pose a safety problem, leave leaf litter and 

mulch, uncultivated soil, dead logs, and snags, 

which are important to many kinds of life from 

native bees to birds.

• Plant and protect a variety of species. Complex, 

multistoried plantings are more attractive to diverse 

animals than plantings of only a few species.

• Create a variety of habitats within the area. Tall 

trees, smaller shrubs, herbaceous plants, and standing dead snags provide important niches where 

different creatures can find their preferred shelter, 

food, and so forth.

• If possible, try to create corridors—connected areas 

where birds and insects can live or travel through 

safely. Ideally, the corridors will be contiguous, 

without large gaps. If the nursery is near a larger 

forest, park, or riparian area, creating a corridor 

between this area and the new planting may help 

species colonize the refuge area. Cooperation with 

neighbors may also help create larger and wider 

corridors. 

Another recommendation is to ensure alternate hosts 

for certain pests are not grown near the nursery. For example, not all rust fungi have alternate hosts but those that do 

can be controlled by removal of these alternate hosts.

Diversity

Nursery areas can sometimes be devoted to a single 

plant species grown closely together—a potential buffet 

for the pests of that species. Studies in silviculture, agroforestry, and sustainable agriculture illustrate the benefits 

of mixed plantings for reducing pest problems. For example, in Southern Florida and the Caribbean, the mahogany 

shoot borer (Hypsipyla grandella [Lepidoptera: Pyralidae]) 

has a major effect on the survival and health of native West 

Indies mahogany seedlings and trees (figure 14.11). To 

reduce the likelihood of attack by mahogany shoot borers, 

researchers and growers are moving away from mahogany 

monocultures and into diversified plantings, with some 

success (Howard and Merida 2010). Reasons for the effec288 Tropical Nursery Manual

Figure 14.10—Sometimes called “the gardener’s best friend,” 

anoles (Anolis species) eat many pest insects including moths, 

locusts, and cockroaches. Photo by Brian F. Daley.
tiveness of diversification range from not attracting large 

numbers of pests to providing habitat for greater biodiversity to balance out pest populations. Nursery growers 

who wish to diversify can mix plants of different species 

throughout the nursery instead of growing all the same 

species in one section (see story about “hiding” seedlings 

from rust in Chapter 20, Discovering Ways to Improve 

Nursery Practices and Plant Quality), although they will 

still want to group mixtures of species according to their 

growth stage and culturing needs. They can also diversify 

areas around the nursery as described in the previous 

section. Clients may also be interested in outplanting in 

diverse patterns instead of large monocultures.

Ongoing Process Evaluation

Assessment and Improved 

Effectiveness of Pest Management 

Approaches. 

Take time to assess and learn from problem prevention 

and holistic pest management practices in your nursery. 

Evaluate the nursery’s daily log seasonally and annually to 

assess the effectiveness of current practices. What is working well? Where is there room for improvement? How 

effective are the sanitation practices? Are any pest trends 

noticeable, such as a decrease in one type of pest but an 

increase in another over time? Are any crops being grown 

that are so susceptible to pests that it may not be costeffective to produce them anymore? Do any instances exist 

where some seedlings are unaffected by a pest while others 

are hit hard? If so, what is the difference? Has the presence 

of beneficial organisms and natural pest predators been 

documented? How can new staff be trained to help scout 

for pests, recognize beneficials, and diagnose problems in 

the nursery? Do any management practices exist that might 

warrant a small experiment in the nursery (see Chapter 20, 

Discovering Ways To Improve Nursery Practices and Plant 

Quality)? What lessons did you learn that you can share 

with other growers? 

Taking time to observe and reflect on pest patterns and 

holistic management processes can make it easier year after 

year to prevent problems and manage them effectively if 

they arise.

Acknowledgment

The authors thank Robert L. James, retired plant pathologist, Forest Service for reviewing this chapter.

Problem Prevention and Holistic Pest Management 289

Figure 14.11—The mahogany shoot 

borer (Hypsipyla grandella) has a major 

effect on the survival and health of native 

West Indies mahogany seedlings. Photos 

by Thomas D. Landis.

A B
References

Cloyd, R.A. 2004. Natural instincts. American Nurseryman. 200: 

38–41.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p.

Francis, J. 2009. Biological disease control—grow your own. 

International Plant Propagators’ Society, Combined Proceedings. 59: 292–295.

Harvey, C.A. 1999. The colonization of agricultural windbreaks 

by forest trees: effects of windbreak connectivity and remnant 

trees. Ecological Applications 10:1762-1773

Hemenway, T. 2009. Bringing in the bees, birds, and other helpful animals. Chapter 7 of Gaia’s garden: a guide to home-scale 

permaculture. 2nd ed. White River Junction, VT: Chelsea Green 

Books: 150–172. Chapter 7. 

Howard, F.W.; Merida, M.A. 2010. Hypsipyla grandella, the 

mahogany shoot borer. Featured Creatures (online publication 

of the University of Florida, Department of Entomology and 

Nematology). http://entnemdept.ufl.edu/creatures/trees/moths/

mahogany_borer-english.htm (July 2013).

Jaenicke, H. 1999. Good tree nursery practices: practical guidelines for research nurseries. International Centre for Research in 

Agroforestry. Nairobi, Kenya: Majestic Printing Works. 93 p.

James, R. 2012. Personal communication. Vancouver, WA: Plant 

Disease Consulting Northwest.

Josiah, S.J.; Allen-Reid, D. 1991. Important nursery insects and 

diseases in Haiti and their management. Information Report 

BC-X-331. Victoria, B.C., Canada: Forestry Canada, Pacific Forestry Centre: 51–59.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1990. 

The container tree nursery manual: volume 5, the biological 

component: nursery pests and mycorrhizae. Agriculture Handbook 674. Washington, DC: U.S. Department of Agriculture, 

Forest Service. 171 p.

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1989. 

The container tree nursery manual: volume 4, seedling nutrition 

and irrigation. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 119 p.

Mizell, R.F., III; Short, D.E. 2009 (revised). Integrated pest management in the commercial ornamental nursery. IFAS Publication ENY-336, 2003, edis.ifas.ufl.edu/IG144. Gainesville, FL: 

University of Florida, Florida Cooperative Extension Service, 

Institute of Food and Agricultural Sciences. 

Stace, P. 1995. Windbreak trees for economic biodiversity: a 

habitat for pests, predators, and crop pollinators. The Sixth 

Conference of the Australasian Council on Tree and Nut Crops, 

Lismore, NSW, Australia, 11–15 September 1995. Lismore, 

NSW, Australia: Australasian Council on Tree and Nut Crops. 

http://www.newcrops.uq.edu.au/acotanc/papers/stace.htm 

(July 2013). 

Unestam, T.; Beyer-Ericson, L.; Strand, M. 1989. Involvement of 

Cylindrocarpon destructans in root death of Pinus sylvestris seedlings: pathogenic behaviour and predisposing factors. Scandinavian Journal of Forest Research. 4(4): 521–535.

Vaughan, M.; Black, S.H. 2008. Native pollinators: how to protect and enhance habitat for native bees. Native Plants Journal. 

9: 80–91.

Wescom, R.W. 1999. Nursery manual for atoll environments. 

SPC/UNDP/AusAID/FAO Pacific Islands Forests and Trees Support Programme, RAS/97/330. Working Paper 9. 53 p.

Wilkinson, K.M.; Elevitch, C.R. 2000. Multipurpose windbreaks: 

design and species for Pacific Island agroforestry. Agroforestry 

Guides for Pacific Islands #4. Holualoa, HI: Permanent Agriculture Resources. http://www.agroforestry.net. (July 2011).

290 Tropical Nursery Manual
Additional Reading

Arentz, F. 1991. Forest nursery diseases in Papua New Guinea. 

Information Report BC-X-331. Victoria, BC, Canada: Forestry 

Canada, Pacific Forestry Centre. pp. 97-99.

de Guzman, E.D.; Militante, E.P.; Lucero, R. 1991. Forest nursery diseases and insects in the Philippines. Information Report 

BC-X-331. Victoria, BC, Canada: Forestry Canada, Pacific Forestry Centre. pp. 101-104. 

Dumroese, R.K.; Wenny, D.L.; Quick, K.E. 1990. Reducing pesticide use without reducing yield. Tree Planters’ Notes. 41(4): 

28–32.

Lopez, R.A.; Duarte, A.; Guerra, C.; Cruz, H.; Triguero, N. 2002. 

Forest nursery pest management in Cuba. In: Dumroese, R.K.; 

Riley, L.E.; Landis, T.D., tech. coords. National proceedings: 

forest and conservation nursery associations—1999, 2000, and 

2001. Proceedings RMRS-P-24. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research 

Station: 213–218.

Mohanan, C. 2000a. Epidemiology and integrated management 

of web blight in bamboo nurseries in India. Proceedings of the 

4th meeting of IUFRO Working Party 7.03.04—diseases and 

insects in forest nurseries. Research Paper 781. Vantaa, Finland: 

Finnish Forest Research Institute: 107–121. 

Mohanan, C. 2000b. Introduction of rootrainer technology in 

forestry—impact on nursery disease management. Proceedings 

of the 4th meeting of IUFRO Working Party 7.03.04—diseases 

and insects in forest nurseries. Research Paper 781. Vantaa, Finland: Finnish Forest Research Institute: 39–47. 

Thakur, M.L. 1993. Pest management in forest nurseries in arid 

and semiarid regions. In: Puri, S.; Khosla, P.K., eds. Nursery 

technology for agroforestry: applications in arid and semiarid 

regions. New York: International Science Publisher: 329–352.

Thornton, I. 1996. A holistic approach to pest management. 

Nursery Management and Production. 12(6): 47–49.

Problem Prevention and Holistic Pest Management 291
292
293

To promote survival and growth following outplanting, nursery stock must first 

undergo proper hardening. Hardening increases plant durability and resistance to 

stress by gradually acclimating plants to field conditions before outplanting. Without proper hardening, plants are likely to suffer from transplant shock, grow poorly, 

or die on the outplanting site. It is important to understand that native plant nurseries are different from ornamental nurseries in that most native plants planted in 

reforestation and restoration projects must endure an outplanting environment in 

which little or no aftercare is provided.

Hardening refers to practices during the nursery cycle that prepare plants for 

the stresses of handling, shipping, outplanting, and field establishment (Longman 

and Wilson 1998, Landis and others 1999). Plant hardiness primarily develops 

internally, although certain external characteristics such as thickening stems and 

reduced succulence in the foliage are indicators of hardiness. This process takes 

time and a common mistake of nursery growers is not to schedule adequate time to 

harden their crops.

To properly harden plants, it is important to consider the Target Plant Concept 

presented in Chapter 3, Defining the Target Plant, and crop planning presented in 

Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records. Using 

knowledge of the expected conditions of a given outplanting site, nursery cultivation may be adjusted to acclimatize plants for site conditions by promoting specific 

traits. For instance, on sites where drought is anticipated, a larger proportion of 

roots relative to shoots may be desirable to improve plant resistance to moisture 

stress. Based on the client’s anticipated outplanting date, the propagation schedule 

can then be developed to include an appropriate hardening period in the nursery 

before plants are delivered to the client.

In this chapter, we illustrate the importance of proper hardiness in promoting 

plant performance following outplanting, discuss how hardiness changes through 

the nursery growing cycle, describe how plants may be conditioned to prepare them 

for the characteristics of a particular outplanting site, and suggest treatments that 

may be used in tropical plant nurseries to help promote hardiness.

Hardening

Douglass F. Jacobs, Thomas D. Landis, and Kim M. Wilkinson 15

Facing Page: Well-hardened crops at Native Nursery on Maui. Photo by Diane L. Haase.
The Importance of Hardening

As described in Chapter 3, Defining the Target Plant, 

the success of nursery plants is not determined by how 

nice the plants look in the nursery, but rather by how 

well the plants survive and thrive after outplanting. As 

described in Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records, plants need to be cultured 

through three phases of growth: establishment, rapid 

growth, and hardening (figure 15.1). During the establishment and rapid growth phases, ideal conditions for 

development and growth are provided (figure 15.2A). 

These first two growth phases provide plants with optimal levels of all potentially limiting factors (water, light, 

nutrition, and so on) while minimizing environmental 

stresses. Plants should not go out to the field while they 

are still in their rapid growth phase; these plants look lush 

and healthy but are not prepared for the stresses of outplanting (figure 15.2B). The hardening phase of nursery 

culture is essential to acclimate plants to the stresses of 

handling and transport, and the conditions of the outplanting site.

All good growers will do their best to reduce stresses to 

plants when they leave the nursery, as described in Chapter 16, Harvesting and Shipping, and Chapter 17, Outplanting. Nevertheless, nursery plants will be exposed to a 

series of unavoidable stresses when it is time for them to 

leave the nursery. The harvesting process requires moving 

and handling plants, which creates potential for physical 

and internal damage. Following harvesting, nursery plants 

are transported to the outplanting site (figure 15.3). During transportation and handling at the site, plants are often 

exposed to unfavorable environmental conditions until 

294 Tropical Nursery Manual

Figure 15.1—Nursery plants go through three stages of growth: establishment, rapid growth, and hardening. The hardening phase 

is crucial to plant survival after outplanting, yet is too often neglected by nursery managers. Illustration adapted from Dumroese 

and others 2008.

Figure 15.2—Succulent shoot tissue (A) would be damaged 

by the stresses of handling, transport, and outplanting. Plants 

in their rapid growth phase look lush but are not prepared for 

the stresses of outplanting. The hardening phase creates a controlled amount of stress for plants, gradually acclimating them 

to the conditions of the outplanting site (B). Photo A by Douglass F. Jacobs, and photo B by J.B. Friday.

B

A
being outplanted. For instance, a ride in the back of a truck, 

even when protected under a cover, will be bumpy and possibly hot. Sunny and windy conditions on the outplanting 

site can result in overheating or desiccation damage.

The hardening phase creates a controlled amount of 

stress for the plants while they are still in the nursery, 

helping the plants to survive the stresses of handling and 

shipping and acclimating them to the conditions of the 

outplanting site (figure 15.2B). Often, these conditions 

include full sun, low amounts of available nutrients, and 

limited soil moisture. After outplanting (figure 15.4), the 

plants may undergo a period of “transplant shock”. This 

shock is primarily because of moisture stress and lasts 

until the roots are able to grow out into the surrounding 

soil to access water and nutrients, compete with other 

plants, resist insect and animal browse damage, and 

endure extreme temperatures.

Proper hardening takes time, and it is a common mistake to rush the process. This mistake often happens when 

growing more than one crop per season or when growers try to force a little extra height growth with crops that 

grow more slowly than expected. Improper hardening can 

also be an issue if the outplanting window was not properly defined, as described in Chapter 17, Outplanting. For 

example, for areas with a pronounced dry season, the late 

or early arrival of the wet season may complicate hardening schedules. Hardening requirements vary by species and 

outplanting sites, but for many tropical species, hardening 

phases of 4 to 12 weeks are common.

Objectives of the Hardening 

Phase

The objectives of the hardening phase will vary by species 

and outplanting environments. In general the objectives are 

to—

• Slow shoot growth.

• Encourage root and stem diameter growth (for good 

shoot-to-root balance).

• Acclimate to the outplanting environment.

• Condition to endure stress.

• Fortify for survival after outplanting.

Practices to reach these objectives may include—

• Introducing gradual, moderate moisture stress.

• Progressively exposing plants to sun equivalent to 

outplanting conditions (full sun in most cases, partial 

shade for understory plantings).

• Reducing fertilization rates and changing mineral 

nutrient ratios.

• Providing good airflow and wind exposure.

• Culturing for root health and proper shoot-to-root 

balance.

Hardening 295

Figure 15.3—Hardening prepares nursery plants for a series 

of unavoidable stresses they will experience when it is time for 

them to leave the nursery. These stresses include handling and 

transportation, such as a ride in the trunk of a car. Photo by 

Thomas D. Landis.

Figure 15.4—Recently outplanted nursery stock must rapidly 

develop new roots that can grow out into the surrounding soil to 

access water and nutrients, compete with other plants, resist insect and animal browse damage, and endure extreme temperatures. Photo by Douglass F. Jacobs.
Shoot-to-Root Balance

Shoot-to-shoot balance is the ratio of shoot biomass 

to root biomass, not shoot length to root length. It is one 

important way to describe plant size and balance. Growing nursery plants to the appropriate size for a specific 

container size is critical. Plants grown too long in small 

containers or too close to one another become tall and 

spindly and do not have enough stem strength to resist 

physical stresses after outplanting (figures 15.5A, 15.5B). 

In addition, these “top-heavy” plants do not have enough 

roots to provide moisture to the foliage, so water stress 

can develop after outplanting. In tropical areas, topheavy plants face the additional risk of storm-force winds 

and are susceptible to blow overs, particularly in the first 

few years after outplanting. Roots in containers that are 

too small often begin to spiral and become compacted 

(figure 15.5C). In these “rootbound” plants, most roots 

become woody and less effective in water uptake and, 

after outplanting, do not grow out from the compacted 

root mass to promote structural stability.

One key to developing a plant with a sturdy shoot and 

well-balanced root system is to select a container that is 

appropriate for the species and conditions on the outplanting site (figure 15.6). Plants should be moved from 

the shaded or protected areas in the nursery as soon as 

they have reached their target height. Experienced growers know that moving plants from a protected area of the 

nursery to an open compound is an easy and effective 

way to keep them in proper shoot-to-root balance. Managing light, nutrition, airflow, and other factors are also 

important, as described in the following sections.

Conditioning Plants for 

Outplanting

To induce plant hardiness and properly condition plants 

to resist stresses, nursery practices are gradually adjusted. 

296 Tropical Nursery Manual

Figure 15.5—This native plant grew way too large for such a small and inappropriate container; it is unlikely that this tree would flourish 

if outplanted (A). These tall, spindly seedlings were grown close to one another and have been held too long (B). These seedlings are rootbound from being held too long (C). Photo A by J.B. Friday, photo B by Diane L. Haase, and photo C by Thomas D. Landis.

Figure 15.6—Some keys to developing a plant with a sturdy 

shoot and well-balanced root system are to select an appropriate size container; to move plants from protected areas to open 

compounds; and to otherwise manage light, nutrition, airflow, 

and water. Photo by Ronald Overton.

A B C
These adjustments must not be too severe, however, because 

overly stressed plants will actually be less hardy. To understand how nursery practices affect hardening, growers need 

to know the role that environmental conditions play in creating hardiness in plants. In the tropics, the main environmental factors that affect plant hardiness are light, water, 

and nutrition (figure 15.7). Wind, airflow, and other conditions also affect plant hardiness. When nursery plants have 

reached their optimum (“target”) size, growers adjust shade/

sunlight, water stress, fertility, airflow/wind, and physical 

factors such as root or shoot pruning to slow shoot growth 

and induce hardiness.

In determining how to properly condition plants for 

the intended outplanting site, it is important to consider the characteristics of the species and the outplanting site. For instance, is this a light-demanding species 

or a shade-loving species? Will plants be outplanted in 

an open field or underneath an existing canopy of trees? 

Will the site be prone to extended dry periods? Do other 

extreme site conditions exist such as high wind, poor 

soils, or salt spray? Understanding the character of the 

site is best accomplished by interacting closely with the 

client ordering the plants. These factors all reflect the 

main principles of Chapter 3, Defining the Target Plant, 

which suggest that the characteristics of nursery stock be 

matched to those of the intended outplanting site. Experience is the best teacher—experiment on a few plants 

and discover which hardening practices work best in 

your nursery circumstances and outplanting sites.

Water

Reducing irrigation duration or frequency creates a 

mild moisture stress, slows shoot growth, and helps condition nursery stock to withstand drier conditions on 

outplanting sites (figure 15.8A). This reduced water availability decreases the possibility of producing top-heavy 

plants and encourages the formation of smaller leaves 

with thicker cuticles that transpire less (lose less water) 

after outplanting. Smaller shoots are also less likely to be 

physically damaged during transplant.

Hardening 297

Figure 15.7—Nurseries manipulate environmental factors—light, 

nutrition, airflow, and water—to slow shoot growth and induce 

hardiness (A). At the Metropolitan Arboretum nursery for Parque 

Doña Inés in Puerto Rico, native palm seedlings are hardened on 

weed barrier cloth in full sun (B). The pots are mulched with locally 

available white stones to reduce weeds and water loss from evaporation. Illustration A by Jim Marin, and photo B by Brian F. Daley.

Figure 15.8—Reducing irrigation to induce a mild moisture 

stress helps harden crops (A). Severe water stress to the point that 

plants wilt (B) is harmful, however. Photos by Thomas D. Landis.

B

A

B

A
Watering frequency needs to gradually be reduced to 

ensure that plants do not permanently wilt or experience severe water stress (figure 15.8B). Adjusting irrigation for hardening requires close observation and 

experience. The best way to quickly and accurately 

evaluate the water status of container plants is to weigh 

the growth container. See Chapter 11, Water Quality 

and Irrigation, for a discussion of irrigation monitoring 

with container weight.

After plants are hardened, they will still require a 

full watering before outplanting and will need good soil 

moisture availability at the outplanting site during their 

early establishment in the field. Many tropical areas have 

pronounced wet and dry seasons. Planting shock can 

often be minimized by outplanting after the onset of the 

rainy season, as described in Chapter 3, Defining the Target Plant, and Chapter 17, Outplanting.

Sunlight

The use of sunlight and shading as a conditioning 

treatment depends on the conditions on the outplanting 

site and the light needs of the species. Plants that will be 

planted into full sun conditions should receive minimal 

or no shading during nursery cultivation, especially during the hardening phase. If plants were started in a covered area (figure 15.9A), they need to be progressively 

exposed to a level of sunlight equivalent to the outplanting site (full sun in most cases, partial shade for understory plantings) (figure 15.9B). Growing plants in outdoor areas also exposes them to ambient temperatures.

Install shadecloth or move the crop to a shadehouse 

to reduce the amount of light a crop receives when it 

will be outplanted in understory or partial shade conditions. Shading is probably an overused treatment in 

nurseries, however, because most species (even those 

classified as shade tolerant) tend to grow best in full sunlight. In addition, many tropical plants often grow excessively in height (“stretch”) under high shade, which may 

create a shoot-to-root imbalance. Nonetheless, if the species is shade loving and will be planted onto a site underneath an existing canopy, then shading may be a useful 

treatment.

Mineral Nutrition

Reducing or stopping fertilization along with reducing irrigation slows shoot growth and hardens plants. 

Among the mineral nutrients, nitrogen, particularly 

in the ammonium form, is the primary driver of shoot 

growth. During hardening, it can be helpful to reduce 

or stop nitrogen fertilization to induce a mild nutrient 

stress. (Note: the use of controlled-release fertilizers with 

long release periods can prevent or delay hardiness from 

developing.)

Some fertilizers have been specifically developed to 

aid in plant hardening, often containing a low-nitrogen–high-potassium formulation. Calcium nitrate is 

also a useful hardening fertilizer because it contains the 

nitrate form of nitrogen, which does not promote shoot 

growth. Calcium also helps develop strong cell walls and 

leaf waxes. Be sure not to use a similar product known as 

298 Tropical Nursery Manual

Figure 15.9—Shade may be used during early phases of plant growth (A), but shade should be removed in the hardening phase to 

expose plants to a level of sunlight equivalent to the outplanting site (B). Photo A by Thomas D. Landis, and photo B by Diane L. Haase.

A B
calcium ammonium nitrate because the ammonium can 

stimulate shoot growth.

Air and Wind

Increased distance between individual containers 

improves air circulation, allows more sunlight to reach 

lower leaves, encourages the development of shorter 

plants with larger stem diameter, and also promotes 

thickening of the leaf cuticle. Containers with individual, 

removable cells can be changed to every other slot to 

increase spacing within the trays during the hardening 

period (figure 15.10).

An interesting wind simulation treatment is known 

as “brushing.” This practice came about after growers 

observed plants repeatedly handled during crop monitoring tended to develop greater stem diameter. The effect is 

replicated by gently moving a horizontal pole (such as a 

clean length of bamboo, or a light PVC pipe) through the 

crowns of the plants in both directions. Of course, this 

practice must be done gently, especially when the foliage is still succulent. Nurseries with traveling irrigation 

booms have mechanized the process by hanging a PVC 

pipe from the boom. A good time to brush plants is right 

after overhead irrigation because the rod also shakes 

excess water from the foliage and reduces the potential 

for foliar diseases such as Botrytis.

Root Culturing

A vigorous, fibrous root system distributed evenly 

throughout the container will rapidly proliferate after 

outplanting. Containers with vertical ribs facilitate 

healthy root structure by limiting root spiraling and are 

designed to promote air pruning at the drainage hole. 

Other root culturing features such as sideslit air pruning 

and copper pruning are effective, especially with vigorous rooted species. See Chapter 7, Containers, for more 

information on these features.

After plants are moved outdoors, it is important not 

to place the containers directly on the ground (figure 

15.11). Instead, plants need to be placed on benches or 

pallets to facilitate air pruning of roots (figure 15.12). 

Otherwise, roots may grow directly into the ground, 

which will require the added task of root pruning during 

harvest. Root pruning immediately before outplanting 

can make the plants more vulnerable to pathogenic fungi 

and may delay quick root outgrowth after outplanting. 

If plants must be on the ground, placing groundcovers 

under them that are impenetrable to roots (such as fabrics treated with copper) can be helpful.

Shoot Pruning

Pruning shoots or “top pruning” is sometimes required 

if the top is too large for the root system. In general, the 

shoots of grasses, forbs, and some shrubs and trees can be 

pruned. However, many trees can be negatively affected 

by shoot pruning, so it is generally only recommended 

as a problem-solving technique for species known to be 

tolerant of pruning. Faster growing plants with a multistemmed form tend to tolerate shoot pruning whereas 

slower growing plants or species characterized by a single 

leading shoot tend not to tolerate shoot pruning. When 

working with new species, a small pruning trial is the best 

way to see how the species responds.

Hardening 299

Figure 15.10—Containers with individual, removable cells can 

be changed to every other slot to increase spacing within the trays 

during the hardening period. Photo by Diane L. Haase.
For species that can tolerate shoot pruning, shoot 

pruning can help maintain a proper shoot-to-root balance 

and reduce water stress resulting from excessively high 

transpirational demand. Pruning also stimulates more 

stem and root growth. Pruning should be done just above 

the height of the smaller plants that have been overtopped 

(figure 15.13). This practice results in additional light for 

smaller plants and helps them reestablish a growth rate 

that is consistent with the rest of the crop.

It is critical that shoot pruning treatments not be too 

severe; a rule of thumb is never to remove more than 

one-third of the total shoot. Pruned plants should always 

be healthy and have enough stored energy to rapidly grow 

new tissue. The International Society of Arboriculture 

300 Tropical Nursery Manual

Figure 15.12—Plants in a hardening area need to be placed on 

benches designed to facilitate air pruning and prevent roots from 

growing into the ground (A) such as the benches used in this nursery 

in Rota (B). Photos by Thomas D. Landis.

Figure 15.11—Plants grown in direct contact with soil (A) may 

grow into the ground (B). Photos by Douglass F. Jacobs.

B

A

B

A
(http://www.isa-arbor.com/) publishes extensively on when 

and how to best prune plants for improved plant health 

and form. It is best to prune succulent tissue because 

woody stem tissue has less regenerative ability.

Some growers have delivered “striplings” for outplanting, 

which are severely pruned saplings, sometimes with foliage 

also removed. The perceived advantage to striplings is they 

are easy to transport, harder to damage in shipping, and 

lose less water to transpiration immediately after planting. This extreme practice is not recommended because 

stripping plants of their foliage often results in mortality 

after outplanting.

References

Landis, T.D.; Tinus, R.W.; Barnett, J.P. 1999. The container tree nursery manual: volume 6, seedling propagation. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 167 p.

Longman, K.A.; Wilson, R.H.F. 1998. Protecting growing trees: preparing young trees for planting. In: Longman, 

K.A. Growing good tropical trees for planting. London, 

United Kingdom: Commonwealth Science Council: 155–158.

Hardening 301

Figure 15.13—For species that can tolerate shoot pruning, the 

objective is to reduce the height of taller plants (A), thereby exposing 

smaller plants to more light and allowing them to “catch up” (B). 

Illustrations by Jim Marin.

B

A
302
303

Plants are ready for harvest and delivery to clients after they have reached target 

specifications (see Chapter 3, Defining the Target Plant) and have been properly 

hardened to withstand the stresses of handling and outplanting (see Chapter 15, 

Hardening). The “harvesting window” is the time period during which plants are at 

target size, maximum hardiness, and most tolerant to stress, that is, they are in the 

best condition for harvesting, shipping, and outplanting. The harvest window timing needs to be coordinated with the client’s outplanting schedule to coincide with 

optimum conditions on the outplanting site.

Tropical plants do not achieve a deep dormancy condition and are therefore difficult to store and can be vulnerable to handling stresses. Plants need to be handled 

with utmost care at all times to minimize effects of temperature, moisture, or physical stresses. It is critical that plants be transported quickly and outplanted as soon 

as possible after leaving the nursery. This chapter describes proper scheduling, harvesting, handling, and shipping procedures for maintaining plant quality from the 

nursery to the field.

Harvesting and Shipping

Diane L. Haase, Thomas D. Landis, and Tara Luna 16

Facing Page: A nursery staff member from the University of the Virgin Islands, Agriculture Experiment Station, delivers a 

healthy black olive (Bucida buceras) tree to a satisfied project partner. Photo by Brian F. Daley.
Scheduling the 

Harvesting Window

In tropical nurseries, scheduling harvesting is based on 

the condition of both the plants and the outplanting site. 

Some tropical climates allow for a wide planting window 

so plants can be outplanted nearly year round if they are 

properly hardened and if soil moisture and temperature 

conditions at the planting site are favorable for survival 

and growth. Other tropical climates have pronounced dry 

seasons, monsoon seasons, or other times when planting 

windows are not open. Nurseries growing for projects with 

limited outplanting windows must schedule crop production carefully to ensure crops are not ready too early or too 

late (see Chapter 4, Crop Planning: Propagation Protocols, 

Schedules, and Records).

Site Conditions

For sites where a distinct dry season exists, scheduling 

harvesting according to the calendar along with experience 

of the nursery and field staff can be quite effective. In general, outplanting is most successful when planting is done 

just after the onset of the rainy season. The dates are selected 

based on past weather records and how well plants harvested 

on those dates have survived and grown after outplanting.

Plant Characteristics

Growers use plant characteristics as indicators to help 

determine when plants are hardy enough to harvest and 

outplant. Hardened tropical seedlings have the following 

characteristics (Hall 2003) (figure 16.1):

• Firm, lignified stems, often brown in color.

• Sturdy, well developed crowns with leaves extending 

more than three-fourths of the length of the stem.

• Vigorous, compact, leathery leaves.

Client Communication

One challenging aspect of scheduling the harvesting 

window is coordinating with the client. Clients sometimes want their plants before the nursery has had adequate time for plants to achieve their target size and hardiness. More often, clients postpone the delivery dates, 

thereby increasing the likelihood of plants becoming 

rootbound or vulnerable to pests. Clear, frequent communication is essential to determine exactly when plants 

are needed for outplanting. It is crucial for the nursery 

staff to educate the client about the importance of timing 

and the consequences of hastening or delaying the harvesting window (reduced plant quality, reduced outplanting 

growth and survival, increased costs for hold over, and so on; 

see Chapter 18, Working With People, and Chapter 4, Crop 

Planning: Propagation Protocols, Schedules, and Records).

Grading Plants for Outplanting

Before being sent for outplanting, plants can be graded 

for size and quality according to established standards, outplanting objectives, or specifications agreed upon with the 

client. “Culls” are plants that do not meet the grading criteria or are damaged or deformed. Sometimes these criteria 

are adjusted during the grading process based on other cull 

and shipping factors that become apparent during the process. Typical grading criteria include size measurements 

such as shoot height and stem diameter at the root collar 

(“caliper”) (figure 16.2). In addition, plants are inspected 

for root plug integrity, physical injury, or disease.

With single-cell containers in trays or racks, the typical 

grading process consists of removing each container from 

the rack, grading the plant within it, and then placing the 

container into either a “shippable” or “cull” rack. For polybag or larger containers, seedlings can be graded individually and sorted into separate areas based on size and quality. 

Plants that meet size and quality standards are considered 

shippable and are counted to establish an accurate inventory. The inventory of shippable plants can then be shared 

with the client. The nursery can also use the inventory as a 

304 Tropical Nursery Manual

Figure 16.1—Properly hardened tropical plants ready for harvest have lignified stems, well developed crowns, and healthy, leathery leaves. Photo by Diane L. Haase.
record of plant sales and for planning future crops (see Chapter 4, Crop Planning: Propagation Protocols, Schedules, and 

Records).

Most nurseries grade their stock as part of the harvesting 

process while others ship ungraded stock to the outplanting 

site where they are graded immediately before outplanting.

Processing Cull Plants

Culls that are damaged or diseased are discarded or, 

better yet, incorporated into a compost pile (figure 16.3A). 

Composted culls can be reused as a soil amendment. For 

woody species, culls can be run through a hammer mill, 

tub grinder, or other machinery to hasten decomposition 

and speed-up the composting process (figure 16.3B). On 

a smaller scale the plants can simply be chopped with 

machetes or other hand tools. For some species, undersized but otherwise healthy plants can be held over for 

additional growth or transplanted into larger containers for future outplanting dates. This approach is common with cultivars that can be outplanted over a large 

geographic area or for threatened or endangered species 

where every plant is valuable.

Short-Term Storage

Plants destined for understory plantings or partially 

shaded sites can be held in a shadehouse (figure 16.4A) 

until they are shipped. Plants that have been hardened 

Harvesting and Shipping 305

Figure 16.2—Common grading standards include shoot height, 

stem diameter at the root collar (“caliper”), and root plug volume 

and integrity. Illustration by Jim Marin. Figure 16.3—Culled seedlings, old growing media, and 

other greenhouse waste can composted (A). Woody materials 

and other green waste can be ground into smaller pieces by 

machinery to speed-up the composting process, as shown here 

at the nursery at Parque Doña Inés in San Juan, Puerto Rico 

(B). Photos by Brian F. Daley.

Figure 16.4—Tropical plants can be held in a shadehouse (A) 

or open area (B) until they are shipped. Photos by Diane L. Haase.

B

A

B

A
for full-sun conditions can be held in an open compound 

(figure 16.4B) because they may lose their conditioning 

if stored too long in the shade. Both structures are typically equipped with a reliable water source, so irrigation 

and fertigation are possible (see Chapter 5, Propagation 

Environments).

Larger containers can be stored in wire racks to keep 

them upright (figure 16.5A) or on pallets (figure 16.5B) 

to stop roots from growing into the ground. To aid in 

drainage, prevent seedling roots from growing into the 

soil, and retard weeds, plants can be placed on a layer 

of pea-sized gravel covered with landscape fabric. Fabric impregnated with copper can also be purchased that 

chemically prunes roots as they emerge from the bottom 

of the containers (figure 16.6).

Packing

Tropical container seedlings are typically shipped in 

their containers. Seedlings in trays or racks can be shipped 

as is while seedlings in individual polybag containers or 

plastic pots can be placed in open-topped boxes or crates to 

minimize toppling and protect against mechanical injury 

(figure 16.7).

Because containers are expensive, nurseries may wish 

to remove plants from the containers at the time of packing. If seedlings are removed from containers, however, 

care must be taken to protect the root plug with plastic 

wrap or some other covering to prevent desiccation. As 

an alternative approach, nurseries may wish to charge a 

deposit or develop some other method to ensure containers are returned to the nursery for reuse.

Plants need to be packed for shipping in a manner that 

encourages air exchange and allows for possible irrigation on the outplanting site. Restricted airflow can trap the 

heat generated by plant respiration and result in damaging 

stresses. After the plants are graded and packed, the final 

step before shipping is to clearly mark each group of plants 

with the species, seedlot, number of plants, and other 

important information.

Shipping

When nursery stock is ready to outplant, it must be transported to the client or outplanting site. The most appropriate delivery method depends on the distance involved, the 

number of plants, and the hardiness of the stock. Most nursery stock in the tropics is delivered by truck or sometimes by 

boat. Nursery plants can be subjected to severe mechanical 

shocks during transport, especially on gravel or dirt roads, 

306 Tropical Nursery Manual

Figure 16.5—Racks or benches can be used to store large container stock (A) and pallets can be used to keep plants off the ground 

(B). Photos by Ronald Overton.

Figure 16.6—Copper-treated fabrics are ideal for ground 

storage because they chemically prevent plant roots from growing 

into the ground. Photo courtesy of Stuewe and Sons, Inc.

B

A
and reducing speed will minimize potential injury (Stjernberg 1997). When shipped by boat, the plants need to be 

packed so that they do not come in contact with sea spray, 

which can severely damage foliage and roots, although the 

damage is usually not visible for several days later.

Containers in racks, pots, or polybags are usually placed 

carefully on the floor or stacked on metal or wooden shelves 

inside the delivery vehicle. Large container stock can be transported in the same racks used at the nursery (figure 16.8).

The risk of injury to nursery stock increases with the 

shipping distance. High temperature is the major risk factor during nursery stock transport. The temperature inside 

the truck, van, trailer, or cargo container needs to be monitored during transit. Delivery vehicles should be aluminum 

or painted white to reflect sunlight and parked in the shade 

during stops and when they reach the outplanting site. 

An insulated truck liner can also be used to protect seedlings from heat (Anonymous 2006). Adding “blue ice” in 

the boxes of small shipments can help keep temperatures 

down although it could increase delivery costs. In some 

areas, refrigerated cargo containers are available; they are 

commonly used to transport produce but can be rented for 

transporting plants. Usually no need exists to turn on the 

refrigeration (it might cool the seedlings too much) because 

the containers are well-insulated and can keep plants cool and 

protected during transport even without the refrigeration on. 

Cargo containers can be left at the nursery and outplanting 

Harvesting and Shipping 307

Figure 16.7—Plants need to be packed for shipping to minimize 

toppling and protect against mechanical injury. Photo A by Ronald 

Overton, and photo B by J.B. Friday.

Figure 16.8—Large container stock can be grown in special racks 

like these at the Forest Service J.H. Stone nursery in Central Point, 

Oregon (A). These plant can then be transported to the field in their 

nursery racks (B). Photos by Thomas D. Landis.

B B

A A
site for a few days to allow time for loading and unloading 

but should be monitored for temperature and kept ventilated with a fan.

If open pickups must be used, then plants need to be protected from wind and sun damage by covering with a reflective tarp. A frame needs to be constructed to suspend the tarp 

above the seedlings so it does not crush the shoots or inhibit 

air circulation. Specially constructed Mylar® tarps with white 

outer and silver inner surfaces are available from reforestation supply companies (figure 16.9A). In operational trials, 

plants under such tarps were as cool as those stored in the 

shade (figure 16.9B). Dark colored tarps, such as army green 

canvas tarps, however, allow plants to heat to damaging levels and should never be used (DeYoe and others 1986).

Regardless of the vehicle used for shipping, air circulation created by spacers (such as wooden boards or foam 

blocks) between racks, pots, and boxes can be used to 

reduce heat buildup and to prevent the load from shifting.

Handling

Nursery plants are in a period of high risk from the time 

they leave the protected environment of the nursery until 

they are outplanted. Proper care when handling nursery 

stock during this vulnerable time is critical to ensure that it 

has the best chance for survival and growth after outplanting. During handling and shipping, nursery stock may be 

exposed to many damaging stresses including desiccation, 

extreme temperatures, or mechanical injuries (table 16.1). 

This period incurs the greatest financial risk because nursery plants have reached their maximum value right before 

shipping (Paterson and others 2001). In fact, handling 

nursery stock is considered a more important factor affecting plant quality than the type of outplanting tool (Adams 

and Patterson 2004).

It is important to emphasize to everyone who will be 

handling nursery stock that nursery plants are alive and 

perishable, and so should be treated with utmost care at 

all times. It is a waste of time and money to produce or 

purchase high-quality plants only to have them die or 

grow poorly after outplanting as a result of unnecessary 

stresses.

Plants are best able to tolerate stress when they are 

not actively growing. Nonhardened, succulent plant tissue is much more vulnerable to stresses. Regular monitoring of plant condition, close supervision of nursery 

and field personnel, periodic testing of plant quality, 

and maintenance of detailed records are essential during 

shipping and handling.

Moisture Stress

Desiccation is the most common stress encountered 

during handling, shipping, transport, and planting, 

and can have a profound effect on survival and growth. 

When exposed for only 5 minutes, seedlings can exhibit 

increasing moisture loss with increasing air temperature 

and wind speed (figure 16.10). A comprehensive evaluation of the various types of stresses affecting plants during handling and outplanting (DeYoe 1986) revealed that 

desiccation of the root system was the most damaging factor and that direct sunlight and high temperatures were 

significant only as they increased moisture stress. Plant 

water potential influences every physiological process and 

at stressful levels can greatly reduce growth even if survival 

is unaffected. These damaging effects can persist for several 

seasons after outplanting. Roots are very vulnerable to desiccation because, unlike leaves and needles, they have no 

308 Tropical Nursery Manual

Figure 16.9—Special tarps protect nursery plants from direct exposure to sun and wind during shipping (A). Research has shown 

that reflective Mylar® tarps provide much better insulation than 

standard green canvas ones (B). Photo A by Thomas D. Landis, and 

illustration B modified from DeYoe and others (1986).

B

A
Process

Potential for Stress

Temperature extremes Desiccation Mechanical injuries Storage molds

Nursery storage High Low None Medium

Handling Medium Medium High None

Shipping Medium Low High None

Onsite storage High High None High

Outplanting High High High None

waxy coating or stomata to protect them from water loss. 

Fine root tips have greater moisture content than woody 

roots and are therefore most susceptible to desiccation; 

if fine roots appear dry, then they are probably already 

damaged or dead.

Roots of container plants are protected somewhat by 

the growing medium, which serves as a reservoir of water 

and nutrients. If the plug becomes too dry, however, desiccation damage can be severe. After roots have dried, 

subsequent growth reductions are inevitable, even when 

shoot water potential recovers (Balneaves and Menzies 

1988). Moisture stress can be avoided by making sure 

plugs are kept moist (but not saturated) throughout their 

journey from nursery to outplanting. Container stock 

should be irrigated 1 to 2 days before harvesting depending on weather conditions (Fancher and others 1986). 

This approach allows the plugs to drain to field capacity; 

saturated media is unhealthy for roots; it increases shipping and handling weight and increases the potential for 

mold development.

Temperature Stress

Either hot or cold temperature extremes can quickly 

reduce plant quality during handling and shipping. Exposure to warm temperatures can damage stock by causing 

moisture stress or heat stress. Plants are alive and respiring and when they are exposed to warm temperatures, 

their respiration adds heat to their environment; this 

condition is particularly serious when air circulation is 

inadequate. Maintaining good air circulation will minimize heat buildup because of plant respiration. Exposure 

to direct sunlight results in a rapid temperature increase 

and can quickly dry out plants. During outplanting, the 

nursery stock usually sits for short periods during packing, shipping, and staging. It is important to have any or 

all of these activities occur in the shade to reduce temperature stresses.

Physical Stresses

Rough handling can result in reduced plant performance after outplanting. Each person involved in the handling and shipping of nursery stock needs to receive training on how to minimize physical stresses. The potential for 

physical damage to nursery stock can come from dropping, 

crushing, vibrating, or simply rough handling. Studies 

have shown that the stress of dropping boxes of seedlings 

reduced root growth potential, decreased height growth, 

increased mortality, and increased fine-root electrolyte 

Harvesting and Shipping 309

Table 16.1—Nursery plants are subjected to a series of potential stresses from harvest through outplanting. Adapted from Landis 

and others (2010).

Figure 16.10—When bareroot conifer nursery stock was exposed for 5 minutes, plant moisture loss increased with higher 

temperatures and wind until plant survival and growth were 

adversely affected. Adapted from Fancher and others (1986).
leakage (Tabbush 1986, Sharpe and others 1990, McKay 

and others 1993). Stjernberg (1996) did a comprehensive 

evaluation of the physical stresses that nursery stock is 

subjected to during transport from the nursery to the outplanting site and found that white spruce seedlings produced 

fewer new roots as the distance the seedlings were dropped 

increased (figure 16.11A). Volume growth of these seedlings 

was still depressed 2 years after outplanting (figure 16.11B).

Accumulated Stresses

Nursery plants are at their maximum quality immediately before they are harvested at the nursery, but they 

then must pass through many hands before being outplanted. Outplanting success is dependent on maintaining plant quality by minimizing stress at each phase of the 

operation. As stress increases, the plant shifts energy from 

growth to damage repair. Physiological functions are damaged and survival and growth are reduced. These effects are 

exacerbated further when plants are outplanted on harsh 

sites. Extreme careless handling of planting stock usually 

manifests itself immediately after outplanting—plants die 

within days or weeks. Stressful injuries incurred between 

harvesting from the nursery and outplanting, however, 

may not always be evident until several weeks or months 

after planting. Symptoms include browning, chlorosis (yellowing), poor survival, or decreased growth.

It is useful to think of plant quality as a chain in which 

each link represents one of the events from harvesting at the 

nursery until planting at the outplanting site (figure 16.12). 

Because all types of abuse or exposure are cumulative, think 

of nursery plant quality as a checking account. Plants are at 

100 percent of quality when they are at the nursery, and all 

stresses are withdrawals from the account (figure 16.13). Note 

310 Tropical Nursery Manual

Figure 16.11—When bags of conifer seedlings were dropped 

from different heights, their ability to produce new roots (root 

growth capacity) was significantly reduced (A). This mechanical injury still affected plant growth 2 years after outplanting 

(B). Adapted from Stjernberg (1996). [Metric conversions: 1 in 

= 2.5 cm; 1 ft = 0.91 m]

Figure 16.12—Nursery plants are subjected to a series of stresses from the time they are harvested to when they are outplanted. Each 

stage in the process represents a link in a chain, and overall plant quality is only as good as the weakest link. Illustration from Landis and 

others (2010).

B

A
that it is impossible to make a deposit; nothing can be done to 

increase plant quality after a plant leaves the nursery. Therefore, care must be taken during all the harvesting and shipping processes to help ensure outplanting success.

References

Adams, J.C.; Patterson, W.B. 2004. Comparison of planting bar and 

hoedad planted seedlings for survival and growth in a controlled environment. Connor, K.F., ed. Proceedings of the 12th 

Biennianl Southern Silvicultural Research Conference. GTR 

SRS-71. Asheville, NC: U.S. Department of Agriculture, Forest 

Service, Southern Research Station: 423–424.

Anonymous. 2006. Greenhouse on wheels: new shipping technology converts dry vans into nursery stock haulers. Digger. 

50: 46–47.

Balneaves, J.M.; Menzies, M.I. 1988. Lifting and handling procedures at Edendale Nursery: effects on survival and growth of 

1/0 Pinus radiata seedlings. New Zealand Journal of Forestry 

Science. 18: 132–134.

DeYoe, D. 1986. Guidelines for handling seeds and seedlings 

to ensure vigorous stock. Special Publication 13. Corvallis, OR: 

Oregon State University, Forest Research Laboratory. 24 p.

DeYoe, D.; Holbo, H.R.; Waddell, K. 1986. Seedling protection 

from heat stress between lifting and planting. Western Journal 

of Applied Forestry. 1(4): 124–126.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual 

for native plants: volume 1, a guide for tribal nurseries. Agriculture Handbook 730. Washington, DC: U.S. Department of 

Agriculture, Forest Service. 302 p.

Fancher, G.A.; Mexal, J.G.; Fisher, J.T. 1986. Planting and handling 

conifer seedlings in New Mexico. New Mexico State University, 

Cooperative Extension Service (NMSU CES), Circ. 526. Las Cruces, 

New Mexico: NMSU CES. l0 p.

Hall, K.C. 2003. Manual on nursery practices. Kingston, Jamaica: 

Ministry of Agriculture, Forestry Department. 69 p.

Landis, T.D.; Dumroese, R.K.; Haase, D.L. 2010. The container

tree nursery manual: volume 7, seedling processing, storage,

and outplanting. Agriculture Handbook 674. Washington, DC: 

U.S. Department of Agriculture, Forest Service. 200 p.

McKay, H.M.; Gardiner, B.A.; Mason, W.L.; Nelson, D.G.; Hollingsworth, M.K. 1993. The gravitational forces generated by 

dropping plants and the response of Sitka spruce seedlings to 

dropping. Canadian Journal of Forest Research. 23: 2443–2451.

Paterson, J.; DeYoe, D.; Millson, S.; Galloway, R. 2001. The handling and planting of seedlings. In: Wagner, R.G.; Colombo, 

S.J., eds. Regenerating the Canadian forest principles and practice for Ontario. Markham, Ontario, Canada: Ontario Ministry 

of Natural Resources and Fitzhenry & Whiteside Ltd.: 325–341.

Sharpe, A.L.; Mason, W.L;, Howes, R.E.J. 1990. Early forest performance of roughly handled Sitka spruce and Douglas-fir of different 

plant types. Scottish Forestry. 44: 257–265.

Stjernberg, E.I. 1996. Seedling transportation: effect of 

mechanical shocks on seedling performance. Tech. Rep. 

TR-114. Pointe-Claire, Quebec, Canada: Forest Engineering 

Research Institute of Canada. 16 p.

Stjernberg, E.I. 1997. Mechanical shock during transportation: effects on seedling performance. New Forests. 13(103): 

401–420.

Tabbush, P.M. 1986. Rough handling, soil temperature, and root 

development in outplanted Sitka spruce and Douglas-fir. Canadian 

Journal of Forest Research. 16: 1385–1388.

Harvesting and Shipping 311

Figure 16.13—It is useful to think of nursery plant quality as 

a checking account in which all types of abuse or stress are withdrawals. Note that all stresses are cumulative and no deposits 

can be made—it is impossible to increase plant quality after 

nursery harvest. Illustration from Landis and others (2010).
312
313

Survival and growth after outplanting are the ultimate tests of nursery plant quality. 

After the nursery plants are established in the field, they will provide many benefits to 

the environment by improving soil quality, enhancing biodiversity, inhibiting establishment of invasive plants, sequestering carbon, restoring native plant populations, 

providing windbreaks, creating wildlife habitat, and preventing soil erosion. In addition, established native and traditional plants can provide food, fuel, medicines, 

crafts, animal fodder, beautification, and many other benefits. Careful planning 

well in advance is important with attention to the eight steps of the Target Plant 

Concept. In addition, care with site preparation, onsite plant handling, selection 

of planting spots, proper planting techniques, support and protection of seedlings, 

and quality control during the outplanting process all help ensure the plants will 

have the best chance to survive. Long-term monitoring by the client and follow 

up with the nursery can help refine target plant requirements and improve future 

project successes.

Outplanting

Diane L. Haase, Thomas D. Landis, and R. Kasten Dumroese 17

Facing Page: Native seedlings ready to be outplanted on a former pasture in Hawai‘i. Photo by Douglass F. Jacobs.
Review of the Target Plant 

Concept and Implications for 

Outplanting

Applying the target plant concept and proper crop 

scheduling are accomplished by working with clients to 

define the target plant for their outplanting site (Chapter 

3, Defining the Target Plant, and Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records). The 

target plant is of the right species and genetic source; is of 

the appropriate size, age, and shape for its purpose; and is 

ready at the right time for outplanting with the best chance 

to thrive on the site. The eight steps for defining the target 

plant are—

Step 1—What are the outplanting objectives?

Step 2—What are the conditions of the outplanting site?

Step 3—What factors on the project site could 

limit success?

Step 4—How will limiting factors be mitigated?

Step 5—What species and genetic sources will 

meet project objectives?

Step 6—What types of plant material (stocktypes) 

are best suited to the project site and objectives?

Step 7—What are the best outplanting tools and 

techniques?

Step 8—What is the best time for outplanting?

Outplanting performance (survival and growth) 

depends on careful consideration of each step. To meet the 

project objectives (step 1), the plant material (steps 5 and 6) 

must be matched to the specific site conditions (steps 2 and 

3). Furthermore, successful outplanting depends on optimum timing (step 8), site preparation (step 4), and proper 

planting techniques (step 7). A quick review of some of 

these key aspects of the target plant as it relates to the outplanting process follows.

The Outplanting Window

The outplanting window is defined as the period of time 

during which environmental conditions on the site are 

most favorable for survival and growth of outplanted nursery stock. Soil moisture is the primary determining factor, 

but other environmental or biological factors can also influence the outplanting window. On tropical sites that have 

wet and dry seasons, the outplanting window needs to be 

as soon as possible after reliable rains begin so that plants 

have sufficient time to develop established root systems 

before the dry season begins. Nursery stock is outplanted 

when soil moisture is high and evapotranspirational losses 

(from wind and sun) are low. Where a very short dry season 

exists, as is the case for the Caribbean side of much of Central America and in some of the wetter Caribbean islands, 

planting can be done throughout the year. To ensure that 

outplanting can commence at the optimum time, careful 

planning is crucial, alongside close coordination with the 

nursery (see Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records).

Limiting Site Conditions

It is critical to identify the outplanting site’s environmental factors that may limit plant survival and growth. 

Temperature and moisture are usually the most limiting 

and are discussed in the following sections. Other site factors, such as aspect and soil type, must also be considered. 

Limiting conditions on each outplanting site must be evaluated well in advance of the actual outplanting to determine 

the optimum planting window.

Soil Moisture

Soil moisture plays a vital role in the uptake and translocation of nutrients and can have a significant influence on 

tropical plant survival and growth (Engelbrecht and Kursar 

2003). Following outplanting, a root system must be able 

to take up sufficient water from the surrounding soil to 

meet the transpirational demands of the shoot. If soil moisture is inadequate, the newly planted seedling can become 

stressed, resulting in lower photosynthetic rates, reduced 

growth, and mortality.

Air Humidity and Wind Speed

Weather conditions at the time of outplanting have a 

direct effect on plant moisture stress. An increase in both 

air temperature and wind speed affect transpiration, especially when relative humidity is low. Relative humidity 

does not influence evapotranspiration rates as much as 

vapor pressure deficit, which is the difference between the 

amount of water the air can hold at a given temperature 

and the amount of water at saturation. Therefore, planting 

is best done during the early morning hours when air temperatures are cool and wind speeds are low. When weather 

is sunny, windy, or dry, it is necessary to take extra protective precautions to minimize plant stresses. In extreme 

cases, the planting operation may have to be suspended 

until conditions are more favorable for outplanting.

Site Aspect and Planting Sequence

Conditions can vary at different locations in the planting area, especially in mountainous terrain. Aspect, or 

314 Tropical Nursery Manual
direction of hill and mountain slopes to solar exposure, can 

have a strong influence on outplanting success. In the northern hemisphere, south- and west-facing aspects have a hotter, drier environment than north and east aspects while in 

the southern hemisphere north- and east-facing aspects have 

a hotter, drier environment than south and west aspects. 

Depending on the severity of moisture or temperature stress, 

shading of outplanted stock can help mitigate these stresses.

Temperature

In tropical regions, low temperatures are only an issue 

for mountainous planting sites and frost is rarely a consideration. Plants destined for these sites need to be adequately 

hardened off to minimize vulnerability to cold damage. 

High temperatures and solar loading are more frequently 

a serious concern during outplanting, especially in lowland 

sites. Ground temperatures in full sun are commonly much 

hotter than ambient temperature. Abandoned pastures 

and other forest restoration/rehabilitation sites are notorious for having extreme ground temperatures that can 

contribute to rapid soil moisture loss and even contraction 

and cracking of the soil. Sun scald or leaf burn on newly 

outplanted plants can result. Even if temperatures are high, 

simple cloud cover can dramatically reduce the plants’ 

solar load and decrease ground temperatures.

Other Limiting Factors

Proper culturing of stock in the nursery can help overcome many other limiting site factors such as lack of beneficial microorganisms and low soil fertility. Other limiting 

site factors can be overcome by site preparation, outplanting methods, and maintenance practices, as described later 

in this chapter.

Species Selection

Choosing the plant species to be outplanted depends on 

the project objectives and needs to be tailored to meet the 

existing conditions at each outplanting site. If the primary 

objective is to restore the site to its natural condition, then 

selection of plant species indigenous to that area could be a 

logical choice. On degraded sites, such as abandoned pastures, however, climatic and edaphic conditions are highly 

altered and vastly different from conditions to which native 

species are adapted. In such cases, it is often more appropriate to plant species (noninvasive) that can withstand the 

extreme conditions. If the site will be managed for agroforestry, then species selection will be based on value for food 

or fodder production. For roadside plantings, trees must be 

able to handle difficult conditions and not produce messy 

flower or fruit that can interfere with cars or pedestrians.

The light requirements of a species are especially 

important to consider. Shade-intolerant species require 

full sunlight and tend to be “pioneer” species that can 

establish following disturbance. Shade-tolerant species can 

survive with less light and tend to grow slower than shadeintolerant species. In addition, some species are capable of 

fixing nitrogen from the atmosphere and can help mitigate 

poor soil conditions. These plant characteristics can be 

useful in deciding which species to use on a specific site. 

For instance, the use of a shade-intolerant, nitrogen-fixing 

pioneer species such as a noninvasive or native Acacia species may be ideal for “badlands,” abandoned bauxite and 

phosphate mine sites, and pasture lands. After those are 

established and site conditions have improved, the site can 

be underplanted with native plant species. In some cases, 

an overstory of exotic trees is established. The canopy of 

these trees functions protectively over the underplanted 

native plant species (figure 17.1), thereby having a nurse 

effect (Feyera and others 2002).

Restoring abandoned pastures is common throughout 

the tropics. These pastures tend to be dominated by exotic, 

invasive species and have unique conditions that require 

special consideration when selecting species for outplanting. Pioneer species generally perform better when planted 

in former pastureland than mature forest species (Holl and 

Aide 2011; Hau and Corlett 2003) and using many species 

is preferable to using few (Van der Putten and others 2000). 

An additional planting at a later successional stage can be 

made to introduce a broader range of species to enhance 

diversity (Lamb and others 2005). Establishing fast-growing shrubs has also been shown to increase tree seedling 

survival (Holl 2002).

The project objectives will also dictate how many species 

to plant on a given site. A single species (“monoculture”) 

Outplanting 315

Figure 17.1—Native plant species underplanted beneath a 

nonnative overstory on Guam badlands. The overstory helped 

to improve the soil, and serves as shelter for the native plants. 

Photo by Ronald Overton.
may be desirable when growing crops or tree farms. The lack 

of diversity in a monoculture may make the site vulnerable to 

disease or insects, but the temporary overstory can stabilize 

the microclimate, provide a forest structure, and encourage 

regeneration (Lugo 1997). Multiple species may be chosen based on their ability to provide multiple benefits or 

to create a multistoried mixture of native trees and shrubs. 

For instance, a mixture of shrubs and trees can provide an 

ideal structure for a windbreak or shelterbelt (Upton and 

deGroot 2008).

For any species used on a site, it is important that they 

meet the target size specifications suited to the particular 

outplanting site conditions (see Chapter 3, Defining the 

Target Plant). For example, smaller plants with shallow 

root systems are best for poorly drained sites. Smaller 

plants also require less water than larger plants. On the 

other hand, larger plants can better compete with existing vegetation. In addition, the proper genetic source 

should be selected (see Chapter 8, Collecting, Processing, 

and Storing Seeds).

Site Preparation

Many tropical sites have become overgrown with invasive plants (figure 17.2) and cannot be restored successfully without intensive site preparation (referred to as 

“site prep”) to achieve adequate control of competing vegetation. Exotic, invasive plants aggressively compete for 

nutrients, water, and sunlight and can severely suppress 

or devastate newly planted nursery plants. In some cases, 

exotic, invasive plants can also pose a severe fire risk.

Removing competing vegetation has several benefits 

(USDA 2002), which can combine to give outplanted 

nursery plants a competitive advantage over exotic, invasive plants. Competitive advantage is critical in tropical 

sites where weed development is rapid and persistent. 

Site preparation is labor intensive and costly, but can be 

well worth the effort. Lowery and others (1993) noted that 

growth and survival of planted trees on tropical sites can 

increase by as much as 50 and 90 percent, respectively, 

with early control of vegetative competition.

Site prep also makes the physical process of planting 

easier by reducing or managing some of the surface debris 

on the site. Some surface debris is a valuable site resource. 

For example, woody material can be chipped and used on 

site as mulch and fallen logs or standing stumps can provide favorable microsites for planting as described later 

in this chapter.

Site prep can be accomplished by mechanical or chemical means. It is important to remember that site prep is 

a form of disturbance and exotic, invasive plants are well 

adapted to successfully invade freshly disturbed sites. 

Therefore, timely replanting of sites where vegetation has 

been removed is important to prevent weeds from recolonizing the cleared area.

Scalping

Scalping is the removal of grasses, forbs, small shrubs, 

and organic debris (“duff”) around planting holes 

(figure 17.3A). Scalping reduces weed competition and 

increases soil moisture availability. Removing organic 

debris around the planting hole also ensures that roots 

are in contact with mineral soil (figure 17.3B); nursery 

stock planted in organic matter or duff tend to dry rapidly and often die. Scalping is ineffective against larger, 

woody, deep-rooted plants.

316 Tropical Nursery Manual

Figure 17.2—Existing vegetation on tropical sites often requires intensive site preparation to reduce competition with 

outplanted nursery stock. Shown: restoration site invaded with 

sword grass (Miscanthus floridulus) and tangantangan (Leucaena leucocephala) in Guam (A); an 8-ft-tall (2.4 m) wall of 

exotic, invasive rubber vine (Cryptostegia grandiflora) smothers 

all remaining woody plants in an abandoned pasture in coastal 

St. Croix, U.S. Virgin Islands (B). Photo A by Ronald Overton, 

and photo B by Brian F. Daley.

B

A
Physical Scalping

Scalping can be accomplished with planting tools such 

as a hoe or the side of a hoedad blade. With other planting 

implements, such as augers, another worker scalps beforehand. Planting contracts often contain specifications for 

the size and depth of scalping. For example, the Forest 

Service requires that all vegetation be removed from an 

area 12 to 24 in (30 to 60 cm) around the planting hole 

and 1 to 2 in (2 to 5 cm) in depth. On exposed sites, duff, 

litter, and decaying plant material should be placed back 

on the cleared surface after planting to serve as mulch 

(USDA 2002). In one study, increasing scalp size resulted 

in significant improvement in growth of planted conifer 

seedlings after 4 years (Rose and Rosner 2005).

Chemical Scalping (Herbicide Application) 

Another site preparation option is to kill competing vegetation on the site with herbicides, especially 

around the planting holes,. Herbicide treatment is particularly effective against grasses and other herbaceous 

plants and is useful in tropical sites where plant growth 

is rapid (Ammondt and others 2013). A general-purpose 

herbicide such as glyphosate is effective against a broad 

spectrum of plant species. Herbicide application temporarily eliminates competition for water and also creates a 

mulch of dead organic matter that reduces surface evaporation. Vegetation control with herbicides has been shown 

to increase subsequent survival and growth of outplanted 

nursery stock. In a study with native tropical hardwoods 

in Costa Rica, tree diameter was significantly greater in 

plots treated with herbicide (Wightman and others 2001). 

An experiment evaluating three levels of vegetation control with chemical scalping significantly increased stem 

volume, basal diameter, and height of conifer seedlings on 

four of five sites with increasing area of weed control, and 

the magnitude of difference between treatments increased 

with time (Rose and Ketchum 2002). Although herbicide 

applications are effective in eradicating nonnative plants, 

they can also kill native plants so judicious and selective 

use is necessary.

The most appropriate herbicide application method 

depends on the type of project. For projects where the 

plants will be planted in rows, herbicides can be sprayed in 

rows by all-terrain vehicles (ATVs). For smaller projects, 

a person trained to select likely planting spots can apply 

herbicides with a backpack sprayer. Where the competing 

grasses and weeds are tall, they are first reduced mechanically, manually, or by burning before herbicides are applied 

to the emerging grasses and weeds. Herbicide application 

can be carefully timed to use the minimum amount possible for the maximum result. Never apply herbicide on the 

same day as planting and always follow the label instructions for application rates and time to reentry to the site 

after application. Quickly filling cleared areas with desirable plants is important to keep weeds from moving back 

in (figure 17.4).

Outplanting 317

Figure 17.3—Scalping is the physical removal of plants and organic debris from around the planting hole (A). Scalping helps to 

control competing vegetation and ensures that plants are planted 

into mineral soil (B). Illustration A by Jim Marin, modified from 

Rose and Haase (2006), and photo B by J.B. Friday.

Figure 17.4—Competing vegetation can be killed with herbicides 

before planting. Quickly filling cleared areas with desirable plants is 

important to keep weeds from moving back in. Photo by J.B. Friday.

B

A
Mounding

In some tropical sites, clay soils and organic matter 

combined with heavy rainfall can result in poor drainage 

and be an impediment to planted nursery stock. Mounding can be used to treat several potentially limiting factors: 

plant competition, poor soil aeration on wet sites, shallow 

soils, and nutrient deficiencies. Sutton (1993) provides 

a thorough discussion of mounding and how it has been 

used worldwide. For our purposes, we define mounding as 

the mechanical excavating and inverting of soil and sod to 

create planting spots that are higher than the existing terrain. With thick duff layers, the resulting mounds consist 

of a mineral soil cap over a double layer of humus (figure 

17.5). The results of mounding have been generally favorable, at least in the short term. A study found that mounding was an effective alternative to herbicides for establishing pedunculate oaks (Quercus robur) on water-logged 

sites (Lof and others 2006). Mounding has been criticized 

from an aesthetic and ecological standpoint and can have 

a negative effect on other forest values such as recreation 

(Lof and others 2006). So, as with all site preparation 

treatments, mounding needs to be carefully evaluated on 

a site-by-site basis and compared with other site preparation options. It is generally only appropriate in areas 

where waterlogging and heavy rainfall are a problem.

Pit Planting (Zaï Holes)

In arid tropical soils, shallow planting pits can be prepared during the dry season (Kaboré and Reij 2004). The 

planting pit (zaï) dimensions vary according to soil type 

(8- to 16-in [20- to 40-cm] diameter and 4- to 8-in [10- to 

20-cm] depth). The excavated soil is ridged around the pit 

to improve its water retention capacity and the pit is filled 

with composted organic matter. After the first rainfall, the 

pit is planted and covered with a layer of soil. The zaï functions to conserve water and reduce soil erosion.

Preparation of Severely Degraded Sites

Many tropical planting sites are established on degraded 

lands, such as abandoned pasture land, old agricultural 

fields, or mine land. On level, well-drained soils with heavy 

textures, mechanical site preparation using tractors with 

disc harrows can be used (Ladrach 1992). On low, wet sites, 

discing can help remove excess water and increase soil aeration. Subsoiling is effective for treating soils compacted 

from years of grazing or farming or on sites that have a 

natural hardpan.

On severely disturbed restoration sites, unusual site 

preparation to create suitable planting may be required. 

After the eruption of Mt. St. Helens in Washington State, 

the restoration of 150,000 acres (60,700 ha) of timberland 

posed some serious challenges (figure 17.6A). Experiments 

showed that seedlings must be planted in mineral soil to 

survive, which required digging through 1 to 2 ft (30 to 60 

318 Tropical Nursery Manual

Figure 17.5—On sites with heavy duff layers or in waterlogged soils, mounding has proven to benefit plant survival and 

growth. Illustration from Landis and others (2010).

Figure 17.6—Some restoration sites require special preparations 

before they can be planted. The blast zone of Mt. St. Helens in Washington State was covered with volcanic ash (A), which had to be dug 

away so that seedlings could be planted in mineral soil (B). Photos 

by Thomas D. Landis.

B

A
cm) of volcanic ash at each planting spot (figure 17.6B).

In many cases, planting sites must undergo major 

stabilization before planting can occur. Because of their 

steep slopes and the erosive power of water, stream 

banks must be stabilized with bioengineering structures 

before they can be revegetated (figure 17.7A). Woody 

cuttings of many riparian species used in the structures 

will sprout (figure 17.7B) and provide rapid revegetation 

(Hoag and Landis 2001). With stabilization, careful planning, and planting quality stock, riparian areas can be 

restored (figure 17.7C).

In other conditions, the disturbance may be so severe 

that recovery has to take place in stages. For example, a 

strategy for addressing watershed restoration, biodiversity, invasive species, and wildfire problems in southern 

Guam involves a diversified landscape approach with 

objectives and prescriptions adapted to local site conditions which range from extremely eroded badlands to 

highly flammable, swordgrass-dominated grasslands. 

Steps include creating fuel breaks, planting nonnative 

Acacia trees to shade out sword grass and rehabilitate 

the soil, and ultimately planting native trees (Bell and 

others 2002).

Onsite Inspection, Storage, and 

Handling

Nursery Stock Inspection

As discussed in Chapter 16, Harvesting and Shipping, 

nursery plants are vulnerable to stresses from the time 

they are harvested at the nursery until they are outplanting. Therefore, it is a good idea to conduct a thorough 

inspection of nursery stock when it arrives at the outplanting site. The plants should appear green and healthy 

with full canopies and without signs of insect, disease, 

or other stresses. Nursery stock should not smell sour 

or sweet, which is evidence that it has been too warm or 

too wet. Root systems need to be moist but not saturated. 

The bark should not easily slough off and the tissue 

underneath should be creamy white, not brown or black. 

If white or gray mycelia (evidence of mold) are present, 

check the firmness of the tissue underneath. Soggy or 

water soaked tissue indicates serious decay and those 

plants need to be culled. Plants with superficial mycelia 

without corresponding decay need to be planted immediately. Fungal molds will not survive after exposure to 

ambient conditions on the site.

Outplanting 319

Figure 17.7—Stream banks often require bioengineering structures (A) for stabilization. When cuttings of certain species are used, 

they can sprout quickly (B). Careful planning and quality planting 

stock help ensure restoration success (C). Illustration A from Hoag 

and Landis (2001), illustration B from Steinfeld and others (2008), 

and photo C by Brian F. Daley.

B

C

A
Onsite Storage

Nursery stock need to be outplanted upon arrival to 

the project site. Therefore, it is always wise to plan ahead. 

Bring only as much stock as can be planted on a given day. 

Weather delays, distance, worker scheduling, poor communication, or other logistical issues can make it operationally impossible to complete planting in a single day, 

however, and therefore short-term onsite storage may be 

needed. The duration of onsite storage should last for only 

a day or two.

Whether for a couple hours or a couple days, overheating and desiccation are the major stresses that can 

occur during onsite storage. To dissipate heat and promote good air exchange, the plants need to be set upright 

and placed in a shady area as soon as they arrive on site. 

Plants should never be left in a closed vehicle. Trees and 

other natural shade are often absent on many restoration 

sites, but even when natural shade is available, it can be 

difficult to keep plants in the shade all day. Therefore, 

plan on erecting some type of artificial shade. Tarps or 

shadecloth suspended between poles is effective. Lightcolored tarps are preferred. As shown in Figure 16.9B, 

dark-colored tarps absorb and reradiate solar heat 

(Emmingham and others 2002) and need to be suspended 

above the nursery stock for good air circulation. 

Plant respiration and transpiration rates are a function 

of temperature, which is influenced by sunlight intensity. 

Therefore, check that root plugs are kept moist and plants 

are not under any moisture stress. Irrigating plants on the 

project site is not commonly done but can increase survival 

when site conditions may dry stock. So, the best onsite storage has access to a reliable water source.

On mountainous sites, the plants need to be covered to 

protect the roots from excessive cold if frost is anticipated 

during onsite storage.

Plant Handling

During the planting process, plants should always be 

handled with utmost care. Plants must be protected at 

all times from desiccation, temperature extremes, and 

physical damage. The same principles of careful handling 

described for harvesting and shipping (see Chapter 16, 

Harvesting and Shipping) need to be applied to the planting 

operation. Crews need to be instructed to never toss or drop 

plants. Research shows that dropping seedlings from various heights can result in growth reductions after planting 

(Sharpe and others 1990, McKay and others 1993). Larger 

plants need to be carried by the container or the rootball, 

never by their stems. Planters should never shake or beat 

plants to remove excess growing medium. Each planter 

should carry only as many plants as can be planted in 1 or 2 

hours. On larger reforestation and restoration projects, it is 

most efficient to use runners that carry batches of nursery 

stock from onsite storage to planters.

All planters need to be thoroughly trained in gentle handling and planting procedures. Even high-quality nursery 

stock will die if improperly outplanted. Training is particularly important with volunteers or other inexperienced 

planters. Many of these people lack the skill or strength necessary to properly plant on wildland sites. One option is to 

have an experienced planter create the planting holes just 

ahead of the volunteer crew, and let the volunteers place and 

tamp plants into position. This technique ensures that the 

experienced planter chooses the proper planting spot, creates the desired planting pattern, and makes certain that the 

planting hole is large and deep enough so that plants can be 

situated properly.

Selecting Planting Spots

Pattern and Spacing

The selected pattern and spacing of outplanted plants 

reflects the project objectives. Most planting projects 

specify a desired number of established plants per area 

(table 17.1). These density targets need to be considered 

general guidelines and should never override the selection of planting spots in biologically suitable areas. Trees 

grown for timber or fruit production are often planted 

with uniform spacing in rows (figure 17.8A). Trees 

planted for windbreaks are planted in staggered rows 

positioned perpendicular to the prevailing winds. Where 

ecological restoration is the objective, however, planting 

distribution and spacing will be more representative of 

natural vegetation patterns based on reference sites (see 

Chapter 3, Defining the Target Plant). For aesthetics and 

a more natural look, other projects may choose random 

outplanting of individual plants (figure 17.8B) or outplanting in random groups (figure 17.8C). 

When planting with larger tree species, proper spacing 

is of utmost importance to minimize competition after the 

trees reach maturity. This spacing is generally achieved 

by assigning minimum distances between plants (table 

17.1). It can be advantageous to plant at a higher density 

than the final target density to avoid the need to replant 

after some plants die. In fact, denser planting can promote plant growth by forming a thicket to protect against 

animals, wind, and competing vegetation and by promoting mycorrhizal development (Upton and deGroot 2008). 

If necessary, plants can be thinned out as they mature.

320 Tropical Nursery Manual
Microsites

Choosing the best planting spots is critical and more 

important than exact spacing. Planting in favorable microsites protects nursery stock and greatly improves the probability of survival. Examples of unfavorable planting spots 

include depressions with standing water, rocky spots, deep 

duff, and compacted soils. Plants shaded by a stump, log, 

or large rock tend to grow well, especially on hot, dry sites 

(figures 17.9A, 17.9B). High sunlight on plant foliage causes 

moisture stress and direct sunlight can cause ground temperatures lethal to the seedling. Planting around physical 

Outplanting 321

Figure 17.9—On sites with uneven terrain with physical obstructions, the best planting spots are in microsites in the shade 

of stumps (A) or other debris (B). Compass designations in A and 

B are for the northern hemisphere; planting orientation is the 

opposite for the southern hemisphere. Specific planting spots are 

also prescribed on sites that have been prepared by discing (C) 

or mounding (D). Illustrations A and B adapted from Rose and 

Haase (2006), and illustrations C and D adapted from Heiskanen 

and Viiri (2005).

Figure 17.8—In addition to target plant specifications, the objectives of the outplanting project also affect planting patterns. If 

the objective is timber or agroforestry, then plants are often regularly spaced (A). Most restoration projects are spaced in a more 

random pattern to mimic natural conditions (B) or in a random 

clumped pattern where different species are planted in groups (C). 

Illustrations adapted from Landis and others (2010).

B

B

C

C

D

A

A
obstructions also provides protection from animal damage. Where planting sites have been mechanically prepared 

with disc scarifiers, nursery stock need to be planted on the 

side of the hole in mineral soil (figure 17.9C). On mounds, 

the best planting spot is on the top (figure 17.9D).

Proper Planting Technique

Planting Hole

Good root-to-soil contact is necessary for nursery 

stock to become established on the site and be able to 

readily access water and mineral nutrients. The planting hole needs to be made deep enough so that, when 

filled, the soil is up to the plant’s root collar. For container plants, the root plug needs to be completely covered with mineral soil (figure 17.10A). “J-rooting” and 

unnecessary exposure of the root plug are avoided when 

the planting hole is sufficiently deep (figure 17.10B), but 

the plant should not be planted too deep (figure 17.10C). 

Burying foliage should be avoided. According to Forest 

Service specifications, the minimum-size hole for container stock is 1 in (2.5 cm) deeper than the plug length, 

and at least 3 in (7 cm) wider than the plug at top of the 

hole and 1 in (2 cm) wider at bottom (USDA 2002). By 

making the hole larger than the root system, the planter 

can break up any compacted soil around the root system, 

thereby creating more favorable soil conditions for roots 

to grow.

The planters need to be instructed to plant at the correct depth and not to pull up on the plant to adjust depth 

or straightness. Plants should not be oriented more than 

30 degrees from the vertical plane (figure 17.10D); this 

configuration seems obvious on level ground, but the 

steeper the slope, the more important this orientation 

becomes. Planting holes need to be backfilled with mineral soil without grass, sticks, or rocks (figure 17.10E). It 

is important to firmly tamp the soil around the root plug 

to remove air pockets (figure 17.10F), but refrain from 

stomping around plants to avoid excessive soil compaction or stem injury.

Planting Tools 

Although the choice of the proper planting tool is important, experienced planters can achieve success with a variety 

of implements and may choose to use a combination of tools 

322 Tropical Nursery Manual

Table 17.1—Plant spacing based on regular grids with resultant densities. Adapted from Cleary and others (1978).

Figure 17.10—Nursery stock should be planted properly (A). Common problems include planting too shallow (B), planting too deep (C), 

not placing vertically (D), filling the hole with debris (E), or allowing 

air pockets around the roots (F). Adapted from Rose and Haase 2006.

B

E

C

F

A

D

Spacing (m) Plants per hectare Plants per acre Spacing (ft)

6.4 by 6.4 247 100 20.9 by 20.9

14.8 by 14.8 494 200 4.5 by 4.5

3.7 by 3.7 741 300 12.0 by 12.0

3.2 by 3.2 988 400 10.4 by 10.4

2.8 by 2.8 1,236 500 9.3 by 9.3

2.6 by 2.6 1,483 600 8.5 by 8.5

2.4 by 2.4 1,730 700 7.9 by 7.9

2.2 by 2.2 1,977 800 7.4 by 7.4

2.1 by 2.1 2,224 900 7.0 by 7.0

2.0 by 2.0 2,471 1,000 6.6 by 6.6
depending on site conditions. Planting failures are often 

more attributable to improper technique or handling 

rather than choice of planting tool (Adams and Patterson 2004). Appropriate planting tools and techniques can 

mean the difference between a live or dead plant, and an 

on-budget or over-budget project (Kloetzel 2004). Handplanting methods provide maximum flexibility in plant 

placement and distribution. A well-trained and experienced hand planter can surpass the planting quality and 

generally match the speed of many automated methods, 

especially over rough terrain. Hand planting is especially 

recommended for placing plants into microsites, and 

when planting a mixture of species or stock types. Motorized auger planting may be useful for larger stock sizes, 

especially on open, flat sites with deep soil and few rocks 

or roots.

Dibbles

Dibbles or dibble sticks were among the first tools 

used to plant small container stock, primarily because 

they are easy to use (figure 17.11A). Dibbles are custommade probes that create a planting hole specific to one 

container type and size. Most designs have one or two 

metal foot pedals for forcing the point into the soil (figure 17.11B). After making the hole, the planter simply 

inserts the container plant and moves to the next hole. 

One drawback is the lack of loose soil to cover the top of 

the plug and prevent possible desiccation of the medium. 

Dibbles are most appropriate for lighter textured upland 

soils or alluvial bottomland soils in wetland restoration 

projects. Dibbles should be avoided on heavier textured 

clay soils because they can compact soil and form a glaze 

around the planting hole that can restrict root egress (figure 17.11C). Hollow dibbles are a recent modification that 

extract a core of soil and therefore reduce soil compaction 

(figure 17.11D). The hollow dibble heads are interchangeable to provide the correct dibble for different container 

sizes (Trent 1999). Commercially produced dibbles are 

available for specific container types and sizes, including 

Ray Leach Cone-tainerTM cells and several cavity sizes of 

Styrofoam® block containers (Kloetzel 2004). The ‘o’o bar 

(Hawaiian name) or oso planting stick (Samoan name) 

is a traditional planting tool similar to a dibble stick and 

is useful for breaking through layers of lava to help tree 

roots reach mineral soil (figure 17.12).

Bars

Planting bars originated with bareroot stock, but this tool 

is also used for smaller container plants. Bars are typically 

cylindrical with a wedge-shaped blade welded on the tip, and 

side pedals to help force the blade into the soil. Like dibbles, 

planting bars require little experience or training. The bar 

is dropped and forced into the ground with the side pedals (figure 17.13A), and by working the bar back and forth 

the planting hole is formed. The nursery plant is vertically 

positioned along one cut face (figure 17.13B), and then the 

hole is closed by reinserting the bar into the soil on the opposite side of the planting hole and rocking the bar back and 

forth (figure 17.13C). The final step is to finish tamping any 

loose soil around the plant to remove any air pockets (figure 

17.13D). Planting bars are often preferred for rocky soils but 

should not be used in heavier textured clays where they cause 

excessive compaction. They are also popular on reforestation 

sites with sandy soils. Planting bars are durable and simple 

to maintain, with only occasional blade sharpening required 

(Kloetzel 2004).

Outplanting 323

Figure 17.11—Dibbles are a useful tool for hand planting 

container nursery stock (A). Because they displace soil to form 

the planting hole (B), compaction can be severe enough to restrict root egress (C). Hollow dibbles are an improvement because they remove a core of soil to create a planting hole (D). 

Photo A by Thomas D. Landis, and illustrations B, C, and D by 

Steve Morrison, modified by Jim Marin.

B

C

A

D
Hoedads

Hoedads are one of the more popular and versatile 

tools for outplanting reforestation and restoration stock. 

Brackets, holding the wooden handle to the desired blade, 

are typically brass for extra weight and penetration, or tin 

alloy (“Tinselite”) for lighter applications. Brackets can be 

found in two blade angle configurations: 100° angle for 

applications on gently sloped or flat areas and 90° angle 

for steep ground planting. It is a good idea to purchase and 

keep handy spare blades, handles, and nuts and bolts with 

matching socket or box wrenches. Blades should also be 

regularly sharpened with a metal file or electric grinding 

wheel (Kloetzel 2004). Hoedads are particularly useful on 

steep sites, and on rocky and compacted sites. They are swung 

much like a pick, and it may take several swings to create a 

proper planting hole. With each swing, the planter lifts up 

and back with the butt of the handle to open the planting hole (figure 17.14A). After a proper hole is opened, 

the planter uses the tip of the hoedad to gently loosen soil 

on the sides of the planting hole to avoid any compaction 

effects. Then, the plant is inserted and positioned to the 

proper depth (figure 17.14B). While holding the plant, the 

planter uses the hoedad blade to backfill the soil around 

the plug (figure 17.14C). Finally, the planter gently tamps 

the soil around the plant (figure 17.14D) and moves to 

the next planting spot. If plant competition is a problem, 

or a planting basin is required, the back and side of the 

hoedad’s blade is a useful scalping tool. Some compaction 

in the planting hole can occur from the backside of the 

hoedad’s blade, but compaction is typically less than with 

other methods. Planting rates vary with container size, 

planter skill, and terrain. Kloetzel (2004) reported that 

beginning planters can install 20 plants per hour while 

324 Tropical Nursery Manual

Figure 17.13—Bars are planting tools that create a planting hole by lateral movement (A). The plant is positioned along one side of the planting hole (B), and soil is backfilled by leverage from the other side (C). Soil 

should be gently compacted around the plant with hand or foot (D). Illustrations by Steve Morrison, modified by Jim Marin.

B

A C

D

Figure 17.12—‘O’o bars or oso digging sticks are 

traditional planting tools useful for digging through 

lava to plant seedlings in mineral soil. Photo by Diane 

L. Haase.
experienced planters may reach up to 100 plants 

per hour, and that on wetland planting projects 

with small stock and favorable soil conditions, 

production reached 240 plants per hour.

Shovels

Although standard garden tile spades can be 

used, professional planters use customized shovels with blades long enough to accommodate large 

containers (figure 17.15A). Wooden handles are 

standard but fiberglass models are lighter, and 

reinforced blades (figure 17.15B) can endure 

the vigorous prying action used to open planting 

holes (figure 17.15C). Although not as difficult 

to learn to use as hoedads, planters need to be 

trained to use tree-planting shovels efficiently. 

After the hole is excavated to the proper size and 

depth, the nursery plant is placed in the hole and 

held in a vertical position (figure 17.15D) while 

the planter backfills around the root plug (figure 

17.15E). Soil amendments, fertilizers, and other 

such in-soil treatments are easily applied with 

planting shovels. When using planting shovels, 

keep some spare handles and footpads on hand, 

along with tools for installing parts and sharpening blades (Kloetzel 2004).

Outplanting 325

Figure 17.14—After several swings with the hoedad to create a deep 

enough planting hole (A), the plant is positioned and held (B) while backfilling with soil (C). The final step is to gently tamp the soil around the plant 

to remove any air pockets (D). Illustrations by Steve Morrison, modified by 

Jim Marin.

B

A C

D

Figure 17.15—Shovels are very 

versatile planting tools and are 

ideal for large and deep container 

plants (A). Specialized shovels have 

reinforced blades (B) that open deep 

planting holes without soil compaction (C). While holding the plant 

vertical in the middle of the hole (D), 

soil is backfilled and tamped around 

the root plug (E). Photos A and B by 

Thomas D. Landis, and illustrations 

C, D, and E by Steve Morrison, modified by Jim Marin.

A B C E

D
Mattocks

The mattock (“talacho”) is a versatile tool used for 

digging and chopping (figure 17.16). It is especially useful for cutting through overstory tree roots when doing 

enrichment plantings under native tropical forests. In 

a comparison of three planting tools used for establishing tropical hardwood seedlings, no difference existed in 

seedling performance attributable to the type of tool but 

workers preferred the talacho because it was easy to use, 

it cut roots even better than a machete, and soil did not 

adhere to it as readily as on the other tools (Mexal and 

others 2005). Planting technique with a mattock is very 

similar to the hoedad (figures 17.14, 17.16) but the sharp 

axe-like end can be used to clear roots and other barriers.

Mechanized Dibbles (Planting Tubes)

Mechanized dibbles create a planting hole by compressing soil to the sides and bottom with a pointed pair of 

hinged jaws. The jaws are switched open with a foot lever, 

and a container plant is dropped through the hollow stem 

tube into the hole (figure 17.17). The Pottiputki planting 

tube is the most popular brand and is available in several 

models with different tube diameters. In some models, the 

planting depth is adjustable, which would be necessary 

for stocktypes with longer plugs. One attractive benefit of 

planting tubes is less worker fatigue because the operator 

does not have to bend over. Planting tubes are popular in 

the Northeastern United States and Canada, although they 

are considered expensive to purchase and maintain. In one 

comparison, planting tubes were as effective as dibbles or 

planting bars (Jones and Alm 1989).

Motorized Augers

Power augers have been used in reforestation for 

decades and are becoming popular for restoration projects 

(figure 17.18A). Augers work best in deep soils without 

too many large rocks or roots, and are the best planting 

326 Tropical Nursery Manual

Figure 17.16—Mattocks (left) are versatile tools useful for digging and chopping through roots. They are similar to the hoedad 

(on the right) but have a sharp axe-like end for clearing roots 

and other barriers. Photo adapted from Forestry Suppliers, Inc., 

(2013) by Jim Marin.

Figure 17.17—Planting tubes have pointed jaws that open the planting hole. The plant is then dropped down into the hole 

through the hollow stem. Original illustration by Steve Morrison, and modified by Jim Marin.
tool to use for larger, taller stocktypes. One concern has 

been compaction or glazing on the sides of augered holes 

that occurs under some soil conditions (Lowman 1999); 

this effect can be minimized by rocking the auger bit 

slightly. A gasoline-powered hand drill can be used with 

auger bits from 1 to 4 in (2.5 to 10.0 cm) diameter, and 

the reversible transmission helps when the bit becomes 

stuck (Trent 1999). Larger augers (10 to 16 in [25 to 40 

cm] in diameter) mounted on a skid steer can also be 

used depending on soil type and terrain (figure 17.18B).

One benefit of auger planting projects is that the 

operator selects the location of planting spots and also 

controls the quality of the planting holes (figure 17.18C). 

One operator can drill enough holes for several planters 

to follow and plant the nursery stock (figure 17.18D). 

When scalping is required, the scalper will select the 

planting spots and create the scalp in advance of the 

auger operator. In some soil types, the operator will have 

to excavate extra mineral soil near each hole to ensure 

proper planting. When possible, it is best to rotate the 

auger operator to reduce fatigue.

Digging auger holes deeper than the depth of the container reduces compaction and can promote downward 

root growth. This approach leaves the planter to support 

the plant at the proper depth in the hole, while filling with 

soil from the bottom up (figures 17.8D, 17.8E). Soil settling 

can be a problem with auger planting so it is a good idea to 

mound soil around the base of the plant. A wide variety of 

augers are commercially available for rent or sale. When 

doing large-scale reforestation or restoration projects, it 

is more cost effective to purchase one. If you are inexperienced with their operation, however, it is a good idea 

to rent first to ensure that you have the correct machine 

for the project. Augers are high maintenance planting 

tools, so have an extra one handy, as well as extra parts 

and bits. Well-organized auger teams can reach production rates ranging from 30 to 70 plants per person/hour 

(Kloetzel 2004). In some parts of Hawai‘i, the auger has 

become the ideal planting tool when volunteers or other 

nonprofessional planters are involved, because the planting rate is 2.5 times that of standard hand tools (Jeffrey 

and Horiuchi 2003).

Outplanting 327

Figure 17.18—Augers are effective planting tools and are available in handheld (A) or tractor-mounted (B). One skilled operator can create planting holes (C) while other workers plant the stock and fill the holes by hand (D and E). Photo A by ©Jack Jeffrey 

Photography, photo B by Brian F. Daley, and illustrations C, D, and E by Steve Morrison, modified by Jim Marin.

B

E

C

A

D
Worker Protection

Planting nursery stock can be strenuous. Encourage workers to wear hard hats, safety glasses, and sturdy 

footwear to protect them from sun, insects, and site hazards. If possible, workers need to rotate with each other 

for carrying, digging, planting, and other tasks. Workers 

need to have plenty of fluids available to drink and need 

to take adequate breaks to avoid exhaustion. The time 

and resources spent on worker protection will be offset 

by potential downtime and workers’ compensation claims 

(Kloetzel 2004).

Treatments at the Time 

of Planting

In addition to site preparation to control competing 

vegetation on tropical reforestation or restoration sites, 

other treatments may be applied to plants at the time of 

outplanting to improve survival and growth. These solutions to potential limiting factors need to be identified during the site evaluation and planning process.

Dipping Roots

The seedling root system or container plug needs 

to be moist when it is placed in the planting hole. The 

practice of dipping plant roots in water or a clay slurry 

to protect them from desiccation during outplanting has 

been used for many years, especially for bareroot stock, 

and has proven beneficial on dry sites. Roots of nursery 

plants dry as they are exposed to the atmosphere during 

harvesting and handling and so it makes sense to rehydrate them or apply a coating to protect them. Simply 

dipping the root systems in a bucket of clean water to 

saturate them before outplanting is beneficial. Wetting 

some of the soil and using it to “muddy-in” the hole with 

wet soil and water also helps ensure that no air pockets 

exist around the plant’s roots.

Many commercial root dips are available and most are 

hydrogels, which can absorb and retain many times their own 

weight in water. Little published research exists on hydrogels 

and the results are mixed (Landis and Haase 2012). In a trial 

with Eucalyptus root plugs dipped in a hydrogel slurry, significantly lower mortality rates emerged 5 months after outplanting compared with the controls. The author attributed 

this outcome to increased soil moisture or contact between 

the root plug and the field soil (Thomas 2008).

Water Catchment

When filling the planting hole, a small bund (embankment) can be made around the base of individual trees or 

blocks of trees to prevent runoff of rainwater and direct it 

to the roots (Upton and deGroot 2008). This method can be 

vital during the dry season when plants are becoming established. The raised bund can be made on the downhill side of 

the tree and a trench or small swale is made on the uphill side 

to encourage water to infiltrate into the root zone.

Protection From Animal Damage

Compared with wild plants, fertilized nursery stock has 

higher levels of mineral nutrients and is therefore preferred 

browse by many animals. In many tropical areas, introduced 

pigs, goats, cattle, sheep, horses, deer, and other animals 

severely damage or destroy plantings by grazing, browsing, 

rooting, and trampling (figure 17.19). If the outplanting area 

is known to have a problem with animal damage, then control measures will be necessary. Physical barriers installed 

immediately after planting such as netting, rigid mesh tubing, bud capping, and fencing can be helpful to protect plants 

long enough for them to grow large enough to resist animal 

damage. Most mesh tubes and netting are designed to biodegrade over time. Even still, periodic maintenance is usually required because shoots can get tangled in the mesh and 

deform the stem. After plants grow out of the netting or tube, 

they are again vulnerable to browsing damage.

Fencing is expensive but in areas with dense animal 

populations, exclusion via fencing is often the most effective method. In Hawai‘i, an intensive restoration program 

to rebuild functional ecosystems in the Auwahi forest of 

Maui includes installation of a 7 ft (2 m) tall fence to exclude 

ungulates, application of a 1- to 2-percent concentration of 

328 Tropical Nursery Manual

Figure 17.19—Animals can cause significant damage to outplanted nursery stock such as the browse damage on this seedling 

planted in Guam. Photo by Ronald Overton.
glyphosate to kill exotic grasses, outplant of quick-growing 

native shrubs, and public involvement (Medeiros and 

vonAllmen 2006). Within 10 years, the results were dramatic and could be seen in satellite photos (figure 17.20). 

Weller and others (2011) concluded that effective conservation of native tropical forests requires ungulate 

exclusion, removal of invasive exotic plant species, and 

proactive restoration programs for native species without natural sources of propagules. In some instances, 

“live fencing,” made of species that grow densely and 

have thorns that will keep animals (and people) away 

from the newly planted nursery stock, can be used. This 

approach requires advanced planning, however, because 

it may take 2 to 3 years before the live fence is sufficiently 

developed.

Chemical repellents are another option to protect 

from animal damage. These repellents are less costly than 

physical barriers but their efficacy can be shorter lived. A 

variety of products are available that have an odor (often 

from predators) or taste that is repugnant to wildlife.

Fertilization

Mineral nutrition is a key component of plant performance after outplanting. Studies have shown a positive 

field response to fertilizer applied at the time of planting or 

incorporated into the growing medium of container seedlings (Haase and others 2006). Fertilizer efficacy, however, 

varies with site conditions (Rose and Ketchum 2002). On 

moisture-limited sites, fertilizer salts can build up to toxic 

levels resulting in a negative effect on survival and growth 

(Jacobs and others 2004). Before applying any fertilizer, 

it is crucial to consider the formulation, application rate, 

placement, solubility/release rate, and existing nutrient 

levels on the site.

Phosphorus is quite often deficient for tree growth in 

tropical soils. Boron can also be deficient in tropical soils, 

particularly in volcanic ash soils and soils of basaltic origin. Applications of these elements are routinely made 

at the time of planting and can produce dramatic growth 

responses on nutrient-deficient tropical soils (Ladrach 

1992). Fertilizer responses are often not realized where 

high competition exists from surrounding vegetation. In 

plots treated with herbicide, fertilizer increased height and 

diameter of tropical hardwood seedlings in Costa Rica by 

19 and 31 percent, respectively, but no response occurred 

from the fertilizer used in manually weeded plots (Wightman and others 2001). When soil pH is too high or too low, 

nutrient availability is reduced; combined applications of 

mulch, lime, and fertilizer to highly acidic soils in Palau 

resulted in dramatic growth responses of planted trees 

(Gavenda and Nemesek 2008, Dendy 2011).

Mulches

Mulching with organic or inorganic materials can reduce 

recurrence of vegetative competition for a longer duration 

than initial site preparation. Mulch mats made from materials such as plastic, fabric, sod, or paper are held in place 

with rocks, branches, or stakes. Mulching can also be accomplished with a thick layer of loose organic matter such as 

corncobs, coconut fiber, pine straw, sawdust, or bark chips 

(figure 17.21). In addition to inhibiting growth of competing 

vegetation, mulch insulates soil from temperature extremes, 

helps maintain soil moisture by reducing surface evaporation, and provides protection against soil erosion. Organic 

mulches also have the benefit of providing nutrients to the 

soil and improving soil structure as they decompose.

Although purchase and installation of mulch materials 

can be costly, mulches can significantly improve plant survival and growth on droughty sites. In a study to examine 

the effects of mulches on the survival and relative growth 

rate of three species in a degraded seasonally dry tropical 

forest, soil water content and sapling growth and survival 

were higher in plots mulched with polyethylene than in 

bare soil plots. Sapling survival under organic mulches 

(alfalfa straw and forest litter) were similar, and lowest in 

bare soil (Barajas-Guzmán and others 2006).

Outplanting 329

Figure 17.20—This intensive restoration project by the Leeward 

Haleakala Watershed Restoration Partnership to rebuild functional 

ecosystems in the Auwahi forest of Maui includes fencing to exclude 

ungulates and can be seen by satellite imagery. Source: “Maui Restoration.” 20°38.627’ N 156° 20.519’ W. Google Earth. March 20, 

2011. Accessed March 2013.
Shelters

Tree shelters (figure 17.22A) can protect plants from 

animal damage and can limit the intensity of UV light and 

drying winds that cause damage by desiccation and sunscald. Engelmann spruce (Picea engelmannii) seedling survival after 11 growing seasons increased from 35 percent to 

78 percent when shelters were installed (Jacobs 2011). Tree 

shelters are available in a variety of sizes and colors (allowing varying amounts of solar radiation to penetrate) and with 

or without venting. Selection of a specific shelter is based on 

expected site conditions and the growth habit of the species. 

In a comparison of ventilated and nonventilated shelters, 

ventilation consistently reduced inside shelter temperatures 

by about 5 °F (2.7 °C) (Swistock and others 1999). Plants kept 

in tall, rigid shelters for a long period of time can become 

spindly (reduced stem diameter relative to height) and 

incapable of standing upright after shelter removal (Burger 

and others 1996). Management considerations for using 

tree shelters need to include the costs of purchase, assembly, installation, and annual maintenance. Nevertheless the 

increased cost may be offset by increased survival, thereby 

reducing the need to replant at a later date when competing 

vegetation is established.

Shelters can also be used to protect newly planted plants 

from frost at high elevations (figure 17.22B).

Shading

Ideally, an outplanting site provides adequate materials such as stumps, logs, rocks, or remnant vegetation to 

provide microsites for planting. It is sometimes useful, 

however, to install artificial shade to protect plants from 

damaging heat. Resistance to heat damage increases 

with plant size as the ability of the plant to shade itself 

increases. Shading only the basal portion of the stem can 

be as effective in preventing heat damage as shading the 

entire stem and some foliage, which can also reduce transpiration (Helgerson 1989). Artificial shade materials 

include cardboard, shingles, rigid shade cloth, or other 

330 Tropical Nursery Manual

Figure 17.21—Mulching can reduce competing vegetation 

around the outplanted seedling. Photo by Ronald Overton.

Figure 17.22—Tree shelters can provide favorable microsites to improve seedling growth and establishment on hot, dry sites (A). Frost 

shelters can be used to help decrease heat loss to the night sky and protect plants from frost at high elevations. At 6,500 ft (1,980 m), the Hakalau Forest National Wildlife Reserve on the Big Island of Hawai‘i is high enough that frost can kill seedlings during the winter months; this frost 

shelter can help keep this ‘akala (Rubus hawaiensis) seedling safe (B). Photo A by Diane L. Haase, and photo B by J.B. Friday.

A B
materials and need to be installed on the “sun” side of 

the seedling (south or southwest side for sites north of 

the equator or north or northeast side for sites south of 

the equator).

Monitoring Planting Quality

The best way to determine if planting has been done correctly is to conduct an inspection during or immediately 

after planting. With contract planting jobs, these inspections 

certify whether the work meets specifications, and the results 

are used to calculate payment. A typical planting inspection 

consists of determination of the number and distribution of 

plants and examination of aboveground and belowground 

planting quality. These quality control checks during outplanting help ensure that each plant was installed properly 

and has the best chance of survival possible.

Determination of the Number and 

Spatial Distribution of Planted Plants

Plots are established to determine whether the correct 

number of plants was installed in a given area, whether 

good planting spots were selected, and whether plants 

were properly spaced. See Survey Types section later in this 

chapter for descriptions of plot establishment.

Aboveground Inspection

A representative sample of plants is examined to check 

the quality of the scalping, stem orientation, and planting 

depth. Planting depth is one of the most critical details to 

check and is usually specified in relation to top of the root 

plug or the root collar.

Belowground Inspection

A hole is dug with a planting shovel alongside the 

planted plant to check for proper root orientation, loose 

soil, air pockets, foreign material in the hole, and so on. 

Begin digging the hole far enough away from the main 

stem (10 in [25 cm]) so that the shovel does not disturb 

the roots. Then, gently clear soil away while digging 

toward the plug so that the plug can be inspected in the 

position it was planted. The plant’s root system must be in 

a vertical plane and not twisted, compressed, or jammed 

and the hole should not contain large rocks, sticks, litter, 

or other foreign debris. Soil needs to be nearly as firm as 

the undisturbed surrounding soil with no air pockets. In 

auger plantings, be sure to check soil firmness near the 

bottom of the holes (USDA 2002).

Monitoring Outplanting 

Performance

Reforestation and restoration outplantings can be an 

expensive investment so it makes sense to conduct surveys to track outplanting success over time. An excellent 

guide on how to evaluate restoration plantings is available in chapter 12 of Steinfeld and others (2008). Quality 

control during or immediately after outplanting ensures 

plants are installed correctly. Longer term monitoring 

provides feedback to the nursery and the client, which 

can lead to improved seedling quality and increased outplanting success in subsequent projects. 

When and What To Monitor

Plots of seedlings should be monitored for growth and 

survival during the first month or two after outplanting and 

again at the end of the first year. Growth can be measured 

as height growth and stem diameter growth (figure 17.23). 

Subsequent checks after 3- and 5-year periods will give a 

good indication of plant growth and survival rates. Survival 

can be expressed in percentages; if the client planted 100 

trees, but after 2 months, 20 were dead, the survival is 80 

percent at that point in time.

Some projects define measurable goals within specified time frames as part of their objectives. These goals are 

sometimes called “success criteria” or “desired future conditions.” For example, a reforestation project might set a 

goal of 400 living trees per acre 2 years after outplanting. 

A native plant project might have a goal to reach a certain 

percentage of native groundcover or species composition 

within the first year after outplanting. Monitoring is then 

conducted to measure if these goals were achieved.

The client and the nursery manager then use this performance information to refine the target plant specifications for 

the next crop. The client may also alter his or her outplanting and management practices to achieve better survival and 

growth based on this information.

Survey Types

Two types of surveys, circular plots and stake rows, have 

traditionally been performed, and each has its own advantages.

Circular Plots

The traditional method to determine planting density is to measure 1/100 acre (40 m2) plots that are evenly 

distributed throughout the planting site. An adequate 

sample is about one plot per acre (2.5 plots per hectare), 

Outplanting 331
with usually no more than 30 plots evenly distributed 

throughout the planted area. A 1/100 acre plot has a radius 

of approximately 12 ft (3.6 m), which is established with a 

center stake and a piece of string or twine cut to this length 

(Londo and Dicke 2006). All planted plants within the plot 

are counted, and their tops examined and measured. The 

root system of the plant closest to the center is excavated to 

evaluate planting technique. Record each plot separately on 

a survey form (figure 17.24) using the examination criteria 

shown in figure 17.10.

Stake Rows

Because it is hard to relocate plants on sites with rapid 

weed growth, 10-plant stake rows can be used to make 

plants easier to find in subsequent evaluations. One row 

plot consists of a starting point that can easily be relocated 

and 10 plants staked along a compass bearing. Height, 

diameter, and plant condition are recorded on the data 

form, along with average spacing between plants. Stake row 

data is typically used to determine survival, growth rates, 

and plants per area (Londo and Dicke 2006).

Sampling Design

Systematic, stratified sampling is often recommended 

because plots are located at standard predetermined distances and are therefore easy to establish and relocate. 

Stratification means that the entire population of plants in 

the outplanting area is subdivided into homogeneous units 

before sampling begins. First, strata of uniform conditions 

are identified, and then sample plots are located systematically within these areas (Pearce 1990). These strata could 

be based on species, nursery of origin, planting crew, or 

any other factor that could introduce serious variation. 

Machine-planted stock on abandoned farmland would 

have less variability because conditions are relatively uniform and planter-to-planter variation is not an issue. In 

contrast, considerable variability exists on hand-planted 

projects in mountainous terrain because of differences in 

aspect, soil, and planting technique.

Number of Plots

The number of plots to establish is generally a function 

of two factors: (1) available resources (time and money), 

332 Tropical Nursery Manual

Figure 17.23—Yard or meter sticks can be used to monitor height growth of planted trees as seen here with Senna polyphylla (A) 

and Cordia rickseckeri (B) in a forest enrichment project in the U.S. Virgin Islands. Photos by Brian F. Daley.

A B
and (2) variability of the attributes that will be measured. 

In calculating an appropriate number of plots, statisticians 

are interested in some measure of variability, such as the 

standard deviation of plant heights in the outplanting. 

For example, if a quick check of plant heights shows great 

variability within the site to be sampled, then more plots 

should be added. On the other hand, if the heights appear 

to be very uniform, then fewer plots will be sufficient. If you 

want statistical significance, more complex calculations are 

available to compute appropriate number of plots using an 

estimate of the variability of the attribute and the degree of 

statistical accuracy desired (Stein 1992). Determining the 

number of plots based on variability is often a judgment 

call but, in most cases, a 1- to 2-percent sampling intensity 

is sufficient (Neumann and Landis 1995).

Post-Planting Maintenance

The most significant threat to success of planted seedlings on tropical sites is competing vegetation and animal 

damage. In addition to preventative measures performed 

by site preparation and treatments at the time of planting, 

post-planting treatments may be required to ensure the 

early growth and survival of seedlings.

Weed Control

Aggressive growth of grasses and weeds makes postplanting vegetation control an absolute necessity for the 

successful establishment of nursery plants on tropical 

project sites. Climbing vines can severely damage young 

trees and must be cut several times a year during the first 

years to keep them from shading, deforming, or even 

toppling young trees. Many field sites are infested with 

exotic grasses, such as the kikuyu grass (Pennisetum clandestinum) from Africa, which was introduced into many 

tropical areas a century ago and has effectively colonized 

expansive areas (figure 17.25). This grass has large roots, 

is a fierce competitor for soil moisture and nutrients, 

and is allelopathic (produces growth inhibitors to other 

Outplanting 333

Figure 17.24—Using a standard survey form will ensure that the same information is collected at each plot. Figure by Jim Marin.
vegetation). Tree growth is severely inhibited if this grass 

is not completely controlled around young trees. Herbicides have been found to be more effective than manual 

weed control because they kill the allelopathic roots as 

well as the tops (Ladrach 1992). In a planting of bluegum 

eucalyptus (Eucalyptus globulus) in Colombia, glyphosate was applied to kikuyu grass in a 3 ft (1 m) diameter 

along with hoed planting scalps at the time of planting 

and again 7 months later. After 2 years, tree volumes 

increased by more than 250 percent by the use of herbicide, compared with trees in plots where weeds were 

controlled by hoeing (Lambeth 1986).

Weed control during the first 2 years after planting helps 

the nursery stock to become established and controls the 

undesired species until they can be outcompeted or shaded 

out. As with site prep, weed maintenance after outplanting can be accomplished by mechanical or chemical means. 

Care must be taken not to damage the planted nursery stock, 

however. Intensive weed management and planting desired 

species eventually will increase the site’s resistance to further 

weed invasion by favoring the growth and establishment of 

the desired species.

Animal Control

As with weed control, continued protection against 

animal damage after outplanting can be pivotal to ensuring the project’s success. Periodic monitoring and maintenance needs to be done for installed mesh tubing to ensure 

that the plant is not tangled in the mesh and that it is still 

providing adequate protection; sometimes the tube can be 

slid upward as the plant grows for continued protection of 

the top. Fences need to be examined regularly and mended 

as needed for continued exclusion of animals. 

Long-Term Followup: Refining 

Target Plant Specifications

Both the nursery manager and the client learn from the 

outplanting experiences and monitoring results. Checking in with clients for the first few years (or longer) after 

outplanting is valuable for improving nursery practices, 

outplanting techniques, and overall project successes. As 

described in Chapter 3, Defining the Target Plant, and 

Chapter 4, Crop Planning: Propagation Protocols, Schedules, and Records, the client and the nursery manager must 

work together from the outset to define target morphological and physiological specifications for the plants based on 

the assessment of site conditions, limiting factors, outplanting windows, and so on. Working together to assess 

what worked and what can be improved helps refine the 

target plant for similar conditions and improve results in 

the future. Chapter 18, Working With People, shows how 

to create clear agreements and responsibilities so the client 

and the nursery manager can enjoy an ongoing cooperative 

relationship. See Chapter 20, Discovering Ways to Improve 

Nursery Practices and Plant Quality, for more information 

on capturing lessons learned from clients and outplanting 

experiences.

334 Tropical Nursery Manual

Figure 17.25—Kikuyu grass has formed a thick, dense cover 

across vast areas in Hawai‘i and must be controlled to enable 

establishment and long-term growth and survival of desirable 

plant species. Photo by Diane L. Haase.
References

Adams, J.C.; Patterson, W.B. 2004. Comparison of planting bar 

and hoedad planted seedlings for survival and growth in a controlled environment. In: Connor, K.F., ed. Proceedings of the 

12th biennial southern silvicultural research conference. Gen. 

Tech. Rep. SRS-71. Asheville, NC: U.S. Department of Agriculture, Forest Service, Southern Research Station: 423–424.

Ammondt, S.A.; Litton, C.M.; Ellsworth, L.M.; Leary, J.K. 2013. 

Restoration of native plant communities in a Hawaiian dry lowland ecosystem dominated by the invasive grass Megathyrsus 

maximus. Applied Vegetation Science. 16: 29‒39.

Barajas-Guzmán, A.E.; Campo, J.; Barradas, V.L. 2006. Soil water, 

nutrient availability and sapling survival under organic and polyethylene mulch in a seasonally dry tropical forest. Plant and Soil. 

287: 347–357.

Bell, F.; Falanruw, M.; Lawrence, B.; Limtiaco, D.; Nelson, D. 

2002. Draft vegetation strategy for southern Guam. Honolulu, 

HI: U.S. Department of Agriculture, Forest Service and Natural 

Resources Conservation Service; Government of Guam Division 

of Forestry. 11 p.

Burger, D.W.; Forister, G.W.; Kiehl, P.A. 1996. Height, caliper 

growth and biomass response of ten shade tree species to tree 

shelters. Journal of Agriculture. 22(4): 161–166.

Cleary, B.D.; Greaves, R.D.; Hermann, R.K. 1978. Regenerating 

Oregon’s Forests. Corvallis, OR: Oregon State University Extension Service. 286 p.

Dendy, J. 2011. Low input methods of forest restoration and 

observations of native birds and flying foxes in savanna habitat 

in the Lake Ngardok Nature Reserve, Palau. MS Thesis, University of Hawai‘i at Hilo.

Emmingham, W.H.; Cleary, B.C.; DeYoe, D.R. 2002. Seedling care 

and handling. In: Oregon State University Extension Service: The 

woodland workbook: forest protection. Corvallis, OR: Oregon State 

University Extension Service. 4p.

Engelbrecht, B.M.J.; Kursar, T.A. 2003. Comparative droughtresistance of seedlings of 28 species of co-occurring tropical 

woody plants. Oecologia. 136: 383–393.

Feyera, S.; Beck, E.; Lüttge, U. 2002. Exotic trees as nurse-trees for 

the regeneration of natural tropical forests. Trees. 16: 245–249.

Forestry Suppliers Inc. 2013. Forestry Suppliers website. http://

www.forestry-suppliers.com/. (March 2013).

Gavenda B.; Nemesek J. 2008. Soil quality and land use changes 

on a humid tropical island-Palau. USDA Natural Resource 

Conservation Service, Pacific Islands Area, Mongmong, Guam, 

USA.

Haase, D.L.; Rose, R.W.; Trobaugh, J. 2006. Field performance of 

three stock sizes of Douglas-fir container seedlings grown with 

slow-release fertilizer in the nursery growing medium. New Forests. 31: 1–24.

Hau, B.C.H.; Corlett, R.T. 2003. Factors affecting the early survival and growth of native tree seedlings planted on a degraded 

hillside grassland in Hong Kong, China. Restoration Ecology. 11: 

483–488.

Heiskanen, J.; Viiri, H. 2005. Effects of mounding on damage by 

the European pine weevil in planted Norway spruce seedlings. 

Northern Journal of Applied Forestry. 22(3): 154–161.

Helgerson, O.T. 1989. Heat damage in tree seedlings and its prevention. New Forests. 3: 333–358.

Hoag, J.C.; Landis, T.D. 2001. Riparian zone restoration: field 

requirements and nursery opportunities. Native Plants Journal. 

2: 30–35.

Holl, K.D. 2002. Effect of shrubs on tree seedling establishment in 

an abandoned tropical pasture. Journal of Ecology. 90: 179–187.

Holl, K.D.; Aide, T.M. 2011. When and where to actively restore 

ecosystems?  Forest Ecology and Management. 261(10): 1558–

1563.

Jacobs, D.F. 2011. Reforestation of a salvage-logged high-elevation clearcut: Engelmann spruce seedling response to tree shelters after eleven growing seasons. Western Journal of Applied 

Forestry. 26:53-56.

Jacobs, D.F.; Rose, R.; Haase, D.L.; Alzugaray, P.O. 2004. Fertilization at planting inhibits root system development and drought 

avoidance of Douglas-fir (Pseudotsuga menziesii) seedlings. 

Annals of Forest Science. 61: 643–651.

Jeffrey, J.; Horiuchi, B. 2003. Tree planting at Hakalau National 

Wildlife Refuge—the right tool for the right stock type. Native 

Plants Journal. 4: 30–31.

Jones, B.; Alm, A.A. 1989. Comparison of planting tools for containerized seedlings: two-year results. Tree Planters’ Notes. 40): 

22–24.

Kaboré, D.; Reij, C. 2004. The emergence and spreading of an 

improved traditional soil and water conservation practice in 

Burkina Faso. Paper 114. IFPRI. Washington, DC: International 

Food Policy Research Institute, Environment and Production 

Technology Division. 43 p.

Kloetzel, S. 2004. Revegetation and restoration planting tools: an 

in-the-field perspective. Native Plants Journal. 5: 34–42.

Outplanting 335
Ladrach, W.E. 1992. Plantation establishment techniques in 

tropical America. Tree Planters’ Notes. 43: 125–132.

Lamb, D.; Erskine, D.P.; Parrotta, A.J. 2005. Restoration of 

degraded tropical forest landscapes. Science. 310: 1628–1632.

Lambeth, C.C. 1986. Grass control with the herbicide Roundup 

increases yield of Eucalyptus globulus in Salinas. Res. Rep. 108. 

Cali, Colombia: Carton de Colombia, S.A. 5 p.

Landis, T.D.; Dumroese, R.K.; Haase, D.L. 2010. The container 

tree nursery manual: volume 7, seedling processing, storage, and 

outplanting. Agriculture Handbook 674. Washington, DC: U.S. 

Department of Agriculture, Forest Service. 200 p.

Landis, T.D.; Haase, D.L. 2012. Applications of hydrogels in the 

nursery and during outplanting. In: Haase, D.L.; Pinto, J.R.; Riley, 

L.E., tech coords. National proceedings: forest and conservation 

nursery associations—2011. Proceedings RMRS-P-68. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky 

Mountain Research Station. 53–58.

Lof, M.; Rydberg, D.; Bolte, A. 2006. Mounding site preparation 

for forest restoration: survival and short term growth response 

in Quercus robur L. seedlings. Forest Ecology and Management. 

232: 19–25.

Londo, A.J.; Dicke, S.G. 2006. Measuring survival and planting 

quality in new pine plantations. Tech. Bull. SREF-FM-001. Athens, GA: University of Georgia, Southern Regional Extension 

Forestry. 5 p.

Lowery, R.F.; Lambeth, C.C.; Endo, M.; Kane, M. 1993. Vegetation management in tropical forest plantations. Canadian Journal of Forest Research. 23: 2006–2014.

Lowman, B. 1999. Tree planting equipment. In: Alden J., ed. Stocking standards and reforestation methods for Alaska. Misc. Pub. 

99-8. Fairbanks, AK: University of Alaska Fairbanks, Agricultural 

and Forestry Experiment Station: 74–77.

Lugo, A.E. 1997. The apparent paradox of reestablishing species 

richness on degraded lands with tree monocultures. Forest Ecology and Management. 99: 9–19.

McKay, H.M.; Gardiner, B.A.; Mason, W.L.; Nelson, D.G.; Hollingsworth, M.K. 1993. The gravitational forces generated by 

dropping plants and the response of Sitka spruce seedlings to 

dropping. Canadian Journal of Forestry Research. 23: 2443–

2451.

Medeiros, A.C.; vonAllmen, E. 2006. Restoration of native 

Hawaiian dryland forest at Auwahi, Maui. Fact Sheet 2006–

3035. Reston, VA: U.S. Department of the Interior, U.S. Geological Survey. 4 p.

Mexal, J.G.; Negreros-Castillo, P.; Rangel, R.A.C.; Moreno, R. 

2005. Evaluation of seedling quality and planting tools for successful establishment of tropical hardwoods. International Plant 

Propagators Society, Combined Proceedings. 55: 524–530.

Neumann, R.W.; Landis, T.D. 1995. Benefits and techniques for 

evaluating outplanting success. In: Landis, T.D.; Cregg, B., tech. 

coords. National proceedings, forest and conservation nursery 

associations. Gen. Tech. Rep. PNW-GTR-365. Portland, OR: U.S. 

Department of Agriculture, Forest Service, Pacific Northwest 

Research Station: 36–43.

Pearce, C. 1990. Monitoring regeneration programs. In: Lavender, D.P.; Parish, R.; Johnson, C.M.; Montgomery, G.; Vyse, A.; 

Willis, R.A.; Winston, D. Regenerating British Columbia’s forests. Vancouver, BC, Canada: University of British Columbia 

Press: 98–116.

Rose, R.; Haase, D.L. 2006. Guide to reforestation in Oregon. Corvallis, OR: Oregon State University, College of Forestry. 48 p.

Rose, R.; Ketchum, J.S. 2002. Interaction of vegetation control and 

fertilization on conifer species across the Pacific Northwest. Canadian Journal of Forest Research. 32: 136–152.

Rose, R.; Rosner, L.S. 2005. Eighth-year response of Douglas-fir 

seedlings to area of weed control and herbaceous versus woody 

weed control. Annals of Forest Science 62: 481-492. 

Sharpe, A.L.; Mason, W.L.; Howes, R.E.J. 1990. Early forest 

performance of roughly handled Sitka spruce and Douglas-fir 

of different plant types. Scottish Forestry. 44: 257–265.

Stein, W.I. 1992. Regeneration surveys and evaluation. In: Hobbs, 

S.D.; Tesch, S.D.; Owston, P.W.; Stewart, R.E.; Tappeiner, J.C.; 

Wells, G.E., eds. Reforestation practices in southwestern Oregon 

and northern California. Corvallis, OR: Oregon State University, 

Forest Research Laboratory: 346–382.

Steinfeld, D.E.; Riley, S.A.; Wilkinson, K.M.; Landis, T.D.; Riley, 

L.E. 2008. Roadside revegetation: an integrated approach to 

establishing native plants. Vancouver, WA: U.S. Department of 

Transportation, Federal Highway Administration. 413 p.

Sutton, R.F. 1993. Mounding site preparation: a review of European and North American experience. New Forests. 7: 151–192.

Swistock, B.R.; Mecum, K.A.; Sharpe, W.E. 1999. Summer temperatures inside ventilated and unventilated brown plastic 

treeshelters in Pennsylvania. Northern Journal of Applied Forestry. 16: 7–10.

Thomas, D.S. 2008. Hydrogel applied to the root plug of subtropical eucalypt seedlings halves transplant death following planting. 

Forest Ecology and Management. 255: 1305–1314.

336 Tropical Nursery Manual
Trent, A. 1999. Improved tree-planting tools. Timber Tech 

Tips 9924-2316-MTDC. Missoula, MT: U.S. Department of 

Agriculture, Forest Service, Technology and Development 

Program. 6 p.

Upton, D.; de Groot, P. 2008. Planting and establishment of tropical trees. Propagation and Planting Manuals. Vol. 5. London, 

United Kingdom: Commonwealth Secretariat. 142 p.

U.S. Department of Agriculture (USDA). 2002. Silvicultural practices handbook (2409.17): chapter 2—reforestation. Missoula, 

MT: U.S. Department of Agriculture, Forest Service. 106 p.

Van der Putten, W.H.; Mortimer, S.R.; Hedlund, K.; Van Dijk, 

C.; Brown, V.K.; Leps, J.; Rodriguez-Barrueco, C.; Roy, J.; Len, 

T.A.D.; Gormsen, D.; Korthals, G.W.; Lavorel, S.; Regina, I.S.; 

Smilauer, P. 2000. Plant species diversity as a driver of early succession in abandoned fields: a multi-site approach. Oecologia. 

124: 91–99.

Weller, S.G.; Cabin, R.J.; Lorence, D.H.; Perlman, S.; Wood, K.; 

Flynn, T.; Sakai, A.K. 2011. Alien plant invasions, introduced 

ungulates, and alternative states in a mesic forest in Hawaii. Restoration Ecology. 19: 671–680.

Wightman, K.E.; Shear, T.; Goldfarb, B.; Haggar, J. 2001. Nursery 

and field establishment techniques to improve seedling growth of 

three Costa Rican hardwoods. New Forests. 22: 75–96.

Outplanting 337
338
339

The ability to produce healthy, vibrant plants is an art, a science, and a learned skill. 

Success, however, depends not only on your ability to produce quality plants, but also 

on your ability to work effectively with people. This chapter provides an overview of 

skillfully managing relationships with diverse groups of people including staff, clients, 

and the community (figure 18.1).

Working With People

Kim M. Wilkinson 18

Facing Page: U.S. Virgin Islands Department of Agriculture nursery on St. Croix. This nursery grows both native tree species and 

fruit and vegetable plants for urban and community forestry. Photo by Ronald Overton.

Figure 18.1—Managing a nursery includes working with the community and relating 

with the public. Friends of Hakalau Forest National Wildlife Refuge raised funds to install 

this new water tank to support the greenhouse work at the refuge. Volunteers from the 

Hawai‘i Nature Center installed the sign. The greenhouse propagates some of the rarest 

plants in the world for outplanting on the Refuge. Photo by Dick Wass, Friends of Hakalau 

Forest National Wildlife Refuge.
Interact Based on the Nursery 

Vision and Objectives

Nurseries are founded on a vision, as described in 

Chapter 2, Planning a Tropical Nursery. The vision 

involves a sense of how the human communities and 

local ecosystems could be in the future as a result of the 

efforts put forth by the nursery today and is a key part 

of relating with other people. Regular attention to the 

vision can help to motivate staff (including yourself), 

keep management on track, attract and retain good clients, relate with other growers and researchers, engage 

with the public, and garner support from the community 

at large. In daily interactions with staff, clients, and the 

community, regular reconnection with the vision serves 

as a guide and a reminder of what needs to be done and 

why. Keep your nursery’s vision, objectives, and mission 

statement in a visible place and refer to them when working with people (figure 18.2).

Working With Yourself

Personalities and management styles vary widely among 

effective nursery managers. Some general characteristics, 

however, are important to good nursery management. A 

clear understanding of and commitment to the vision are 

essential along with practical management attributes. 

Consider cultivating the following attributes in yourself:

• Keen observation skills.

• A flexible management style (scheduling must be 

adaptable to the shifting needs of living, growing 

plants).

• An ability to “think like a plant” (managing plants 

effectively is an art as well as a science; someone who 

has a “feel” for the crops will likely do better as a 

manager than someone who approaches crops strictly 

from a technical perspective).

• A sense of responsibility for plants in the nursery.

• A creative approach to working with land managers, 

staff, and other people.

• Enthusiasm for promoting a culture of learning and 

sharing within the nursery and with the larger community. 

A small nursery usually has one person, the manager, 

who takes care of everything. Even so, it is essential to have 

at least one backup person who also understands crop status, knows basic things that need to be done, and knows 

how to do these things. The backup person can keep the 

nursery running and the plants alive and healthy in case the 

primary person becomes unavailable. In a larger nursery, 

the manager oversees the same tasks but delegates many 

of them to employees. Many individuals contribute their 

task to the process, but nursery management cannot be 

conducted “by committee.” All nurseries, large or small, 

ultimately need one person, the manager, to take responsibility for the crop (figure 18.3).

340 Tropical Nursery Manual

Figure 18.2—Regular reconnection with the nursery’s vision 

serves as a guide and a reminder. This “Forest Team” sign reminds everyone the nursery is about more than the trees. Photo 

by Douglass F. Jacobs.

Figure 18.3—Keen observation skills and the ability to “think 

like a plant” are key attributes of an effective nursery manager. 

Photo by Thomas D. Landis.
Working With Nursery Staff

As the manager, you should help each employee connect with the nursery’s vision. When everyone shares and 

is knowingly working towards the vision, each day will be 

a meaningful day, even when filled with tedious tasks such 

as pulling weeds. Sharing the nursery’s mission and overall 

goals fosters a greater sense of purpose among staff thereby 

elevating morale and avoiding “it-is-just-a-job” attitudes 

(figure 18.4).

Staff need to be trained to observe the crops, detect 

problems, and understand and carry out their direct 

responsibilities. Some education in horticulture is helpful 

to enable employees to “think like a plant” and they can be 

encouraged to stay curious and learn more about the plants 

and their production. Safety training is essential.

Clear communication is core to great employee relations. 

Your daily communication with staff will be to assign roles, 

tasks, and goals. You should encourage dialogue and be open 

to feedback from employees about how they think they can 

work more efficiently. You should also provide feedback 

to staff so they can optimize the value of their work. Brief, 

weekly meetings are valuable for linking the day-to-day 

tasks with the nursery’s vision. Meetings can be scheduled 

on Monday morning to assess plans and prioritize activities for the week and again at the end of the week to evaluate progress and identify potential activities for the coming 

week. Connecting to the vision at the beginning and end of 

the week sets the tone for current and future work.

Creating a Learning Organization

Learning takes place on an individual level and also 

within the nursery as an organization. If a nursery is to grow 

and adapt, it is necessary to develop an organizational culture that facilitates learning and collaboration (figure 18.5).

Adaptive learning within an organization depends on 

a willingness to accept new information and ideas, to build 

bridges and make connections among other sources of knowledge, and to occasionally revisit and question fundamental 

assumptions, even if doing so may mean changing “the way 

things have always been done.” Table 18.1 summarizes some 

ways that organizations may or may not be set up for adaptive learning. Creating a nursery culture that facilitates, rather 

than hinders, learning and adaptation is important.

Working With People 341

Figure 18.4—Each person who works at the nursery may 

choose to state in his or her own words his or her vision for his 

or her work and how it links to the nursery’s mission and objectives. Then, even on days when the task is pulling weeds all day, 

it is still a meaningful day. Photo by Susan Charnley.

Figure 18.5—It is important to develop an organizational culture 

within the nursery that facilitates learning and collaborating. 

Nursery staff in Yap work together to test the irrigation system. 

Photo by George Hernández.

Table 18.1—How organizations treat information. Adapted 

from Westrum (1994).

Pathological 

organization

Bureaucratic 

organization

Learning 

organization

Do not want to know May not find out Actively seek 

information

Messengers are shot Messengers are 

listened to

Messengers are 

trained

Responsibility is 

shirked

Responsibility is 

compartmentalized

Responsibility is 

shared

Bridging is 

discouraged

Bridging is allowed 

but neglected

Bridging is 

rewarded

Failure is punished 

or covered up

Organization is just 

and merciful

Inquiry and 

redirection

New ideas are 

crushed

New ideas present 

problems

New ideas are 

welcomed
Discovering

Daily observation and recordkeeping, maintaining plant 

development records, and updating propagation protocols 

are the foundation for understanding how crops grow and 

develop in your nursery environment. Making discoveries through simple experimentation is often a key aspect 

of successful nursery management. Monthly or seasonal 

staff meetings are useful for determine some of the most 

pressing questions facing the nursery. These questions will 

shape priorities for trials. For example, at the beginning 

stages of nursery development when production levels are 

low, the nursery may decide to try different container types 

to determine the best ones for the crops to be grown at the 

nursery, or to experiment with different seed treatment 

techniques for a new species. Later in nursery development, 

other pressing questions may arise such as: What problems are recurring that might be preventable? What could 

improve efficiency? What could improve crop quality? The 

nursery might decide to test seed sources and assess their 

performance in the nursery and the field to refine target 

plant objectives. Ways to design, execute, and assess these 

experiments are detailed in Chapter 20, Discovering Ways 

to Improve Nursery Practices and Plant Quality.

Training

Training and ongoing education is of great value to you 

and your staff. The more that everyone understands his or 

her work and the effects of his or her activities, the more he 

or she will be able to relate to the crop and improve plant 

quality and nursery efficiency. Attending training sessions 

and conferences and reading pertinent publications are 

important investments in the nursery’s adaptability and 

growth (figure 18.6). Visiting other nurseries and reciprocating by hosting other growers at your nursery is an 

important part of cultivating supportive, informative, sharing relationships (figure 18.7) and are a wonderful opportunity to gain a broader perspective.

Working With Clients

Nurseries take many different forms: retail, wholesale, 

research, nonprofit, community-based, or some combination 

thereof. Clients may be farmers, forest planters, conservation 

area managers, schools, landscapers, nonprofit organizations, gardeners, or government agencies. With increasing 

awareness of the importance of quality plants well-matched 

for conditions on the outplanting site, more clients seek nurseries that can provide those kinds of plants. Working with 

clients involves understanding their target plant needs, contracting, ongoing communications, and following-up after 

outplanting to see how the plants performed in the field.

Defining Target Plant Needs

As discussed in Chapter 3, Defining the Target Plant, 

nursery managers and land managers ideally engage in 

an ongoing discussion about project objectives, reevaluate 

successes and failures observed on the outplanting site, and 

subsequently redefine the target plant to improve future 

plant quality in the nursery and on the outplanting site.

342 Tropical Nursery Manual

Figure 18.6—Training and ongoing education is of great value 

to you and your staff. Here, nursery workers gathered for a training to learn about grafting techniques in Volcanoes National Park, 

on the Big Island of Hawai‘i. Photo by Kim M. Wilkinson.

Figure 18.7—Hosting visits to your nursery and attending 

events at other nurseries is a great way to cultivate information exchange and supportive relationships. Photo by Kim M. 

Wilkinson.
For new nurseries, clients, or projects, a good starting 

point for the Target Plant Concept dialog is to consider 

past experiences on similar sites and determine the target plant needs for the project. For complex, large, or specialized projects, another professional may need to assess 

the site and create a plan including appropriate species 

selection, spacing, and other target plant needs. Examples 

include government-supported projects that warrant a 

Forest Stewardship Plan, a Conservation Plan (such as 

the USDA Natural Resources Conservation Service might 

do for soil conservation or riparian restoration), a Habitat 

Improvement Plan, or commercial Farm Plans. If a new client might require this level of assistance, steer the client to 

the appropriate agency or professional and invite him 

or her to order from you when his or her plan is ready 

and the target plant material requirements are defined. 

Sometimes, agencies may have general plant recommendations, such as appropriate windbreak species, requiring 

you and the client to fill in the remaining “blanks” for the 

target plant, or sometimes you and the client may simply 

need to determine outplanting window and local genetics. 

The example from Chapter 4, Crop Planning: Propagation 

Protocols, Schedules, and Records, of a client and nursery 

working together is revisited here (see textbox) with more 

emphasis on how the communication process works.

Working With Noncontracted Clients

Nurseries that grow on contract may also grow additional seedlings or have overstock to sell to potential 

customers that need plants immediately. If the seedlings 

are genetically appropriate for their site, these customers 

may wish to purchase them for their project. A potential customer occasionally may have the notion that your 

nursery can provide “cheap, all-purpose, ready-made, 

grow-anywhere plants”—we know that does not exist. In 

this circumstance, it is your job to help them learn; by 

requiring the target plant information you are supporting the success of their project, which reflects favorably 

on your nursery. Not everyone can be your client, and 

that is okay. If you are having trouble getting a potential 

client to think through his or her project and define his 

or her target plant needs, be friendly but firm and give 

the client a way to come back: “Our nursery takes pride in 

providing high-quality plant materials suitable for client’s 

needs. I would be happy to help you with your plants after 

you have determined your target plant needs for your outplanting site.”

Working With Contracted Clients

Communication about plant targets is easier when clients 

order plants on contract. Many nurseries grow most of their 

native and traditional plants to order on contract, which 

ensures that plants are matched to specific project objectives and outplanting sites. This approach is the opposite of 

speculative growing, where nurseries produce plants based 

on estimates of future demand. Contract growing involves 

producing plants to a client’s target specifications (figures 

18.8, 18.9). The ecological and economic benefits of this 

approach are plentiful—clients benefit by getting exactly 

the plants they need when they need them, nurseries benefit 

by minimizing waste and maximizing time and resources 

propagating only plants that will be used, and plants and 

the environment benefit from greater success in survival, 

growth, and establishment of locally appropriate species. 

For conservation work and habitat improvement, growing 

locally adapted, genetically appropriate plant materials on 

Working With People 343

Figure 18.8—The process of defining target plant materials is an essential tool for specifying what the nursery will grow for the 

client. Illustration adapted from Landis (2011) by Jim Marin.
contract helps maximize benefits to pollinators and other 

wildlife. When ordering plants on contract, clients need to 

give the nursery sufficient time to produce them to specification for outplanting at the correct time.

If the client is a family or an organization with several 

members, ask the client to assign one sole contact person 

for the order with the nursery. This person should be your 

contact from the time of drafting and signing the contract 

through to delivery of the plants. In turn, the nursery should 

have only one contact person for that client. This one-onone arrangement precludes many potential misunderstandings, streamlines time and energy spent on communication, and helps develop long-term relationships 

and trust. For contract growing, create a clear agreement in writing. In some circumstances, such as growing for neighbors or close community members, a written agreement may seem overly formal; however, it can 

be invaluable for creating clear expectations, enhancing 

communication, demonstrating your professionalism, 

and making the entire process a smooth one for everyone 

concerned. Often your client will be new to purchasing 

plants by contract and will be relieved you have a protocol in place to ensure everything goes smoothly, and the 

client’s investment is protected. Both the client and the 

nursery representatives need to sign the agreement, and 

each need to keep a signed copy. 

The terms of the agreement must include the following—

• Description of the plant materials to be provided 

(such as species, genetics, container type, plant size).

• Anticipated schedule.

• Quantity of plants to be provided.

• Price per unit and the total price for the order.

• Any additional fees that may apply, such as delivery 

charges and refundable container deposits.

• When and how payment will be made.

Crop production may vary from year to year. If possible, 

when the order is placed, agree on a window of time for plant 

delivery that spans a few weeks rather than setting an exact 

date. Based on crop development, the exact date for delivery 

can be determined closer to the tentative delivery date.

344 Tropical Nursery Manual

An Example of Working With a 

Contracted Client

Recall from Chapter 4, Crop Planning: Propagation 

Protocols, Schedules, and Records, that... In March, a 

retired cattle rancher on the Big Island of Hawai‘i called 

your nursery. She desires to plant 500 koa (Acacia koa) 

trees inside a fenced area of former cattle pasture as a 

legacy grove for her grandchildren. Her site, at 2,000 

ft elevation on the leeward side of the island, has been 

in pasture for more than a century. She wants to use a 

dibble to plant trees during the November to February 

rainy season. Because her family will help during their 

Christmas holiday, the ideal delivery date is December 

15. Before planting, she will remove the grass.

Based on this information, you begin a conversation 

with the client to discuss the criteria necessary to define 

the target koa seedlings for her site. This conversation 

requires you to carefully listen to what the client needs 

and wants, and, equally important, to offer your expertise 

and recommendations on what is realistically possible. 

The client may not know much about genetics, so you 

share the importance of local adaptation and where to 

find information about appropriate sources for her trees. 

How the koa will benefit from rhizobia and mycorrhizal 

inoculation may also be new information for the client. 

The optimal outplanting time may be the start of the rainy 

season in November, but if she does not have the family 

labor she needs, the timing will have to be scheduled for 1 

month later. Large seedlings might be ideal for this project because of the competing grass issue, but because of 

the client’s budget and time constraints, you will need to 

discuss smaller stocktype options. She may have initially 

planned to outplant with a dibble, but after a discussion 

about how these tools can cause compaction and restrict 

roots, she decides to use a pick and shovel instead. After 

all eight aspects of the target plant are defined, you are 

both clear about what kinds of plants would best meet the 

outplanting objectives and are ready to create a contract 

that clarifies expectations and prevents any surprises at 

plant delivery time.

Figure 18.9—The nursery manager and the land manager 

meet at the outplanting site to define target plant specifications 

for the project. Photo by Brian F. Daley.
The sample contract in the textbox below is for demonstration purposes only. Contracts at your nursery 

should be tailored to meet your and your client’s needs. It 

is best to consult with a legal expert to ensure the contract 

protects your nursery and conforms to local legal statutes.

Ongoing Communications 

With Clients

Clients appreciate staying informed about their crop’s 

progress. Sending e-mail updates (including digital photos), 

having periodic phone conversations, or inviting clients 

to visit their crop at the nursery is vital to keeping them 

involved and committed to the schedule (figure 18.10).

Working With People 345

Sample Contract With Green’s Tree Nursery

This agreement, entered into this 7th (day) of March (month), 2014 (year), by and between Green’s 

Tree Nursery and Upland Ranch and Forest (hereinafter referred to as “Client”) witnesses as follows:

Whereas Green’s Tree Nursery is organized to provide plant materials for outplanting; whereas Client is interested in purchasing plant materials from Green’s Tree Nursery, it is agreed between the parties as follows:

I. Plant Materials Provided by Green’s Tree Nursery

In time for the planting window December 10–20 Green’s Tree Nursery will provide 500 koa (Acacia 

koa) seedlings of the following specifications: 

• Species—koa (Acacia koa).

• Genetic source—Seeds sourced from minimum 50 parent trees of good form (large, straight-boled) 

from koa forest neighboring Upland Ranch.

• Size and description—Containers: Ray Leach “Stubby” cells; Seedling size: 15 to 30 cm tall, minimum stem

diameter 3.5 mm; Roots: firm and nodulating with rhizobia, inoculated with mycorrhizal fungi (AMF).

• Price—$3.50 per seedling.

II. Fees

Client agrees to pay Green’s Tree Nursery $1,750 for the 500 plants listed above. Payment shall be made 

in the following way: an initial fee of $875 (50% of the total for plant materials) is required to begin propagation, 

with the balance of $875 to be paid before dispatch of the plant materials. Other fees, such as container deposits 

(10 cents per “Stubby” cell) and shipping/delivery charges (TBD based on carrier) if applicable, will be billed separately and are also to be paid in full before dispatch of the plant materials.

If any payment as per the above schedule remains overdue for more than 60 days, Client acknowledges that Green’s 

Tree Nursery may take legal action to collect the overdue amount. In such event, Client will be responsible for all reasonable litigation expenses incurred by Green’s Tree Nursery, including, but not limited to, court costs and attorney fees.

III. General Conditions

Green’s Tree Nursery agrees to use its best efforts to provide the plant materials listed in Section I above.

Client understands and acknowledges that Green’s Tree Nursery shall in no way bear liability for results produced 

in use of plant materials. Green’s Tree Nursery’s maximum liability is limited in amount to the amount paid by Client 

to Green’s Tree Nursery for the purchase of the plant materials under all circumstances and regardless of the nature, 

cause, or extent of any loss.

In the event that Client cancels the order for plant materials in whole or in part, Client agrees to pay the balance 

due for the full amount for plant materials as listed in Section I.

Green’s Tree Nursery reserves the right to prorate or cancel any order, in whole or in part, because of natural disaster, disease, casualty, or other circumstances beyond our control. In the event that Green’s Tree Nursery is unable to 

provide the plant materials listed in Section I above by December 20 , the initial fee paid by Client may be applied 

to another purchase, credited to a future order, or refunded, as requested by Client. In any other event, the initial fee 

is nonrefundable and the entire balance is due.

Continued on next page
346 Tropical Nursery Manual

Client agrees that all plant materials ordered must be dispatched (picked up, shipped, or delivered) within 30 days 

of notification of readiness as determined by Green’s Tree Nursery. Plant materials not dispatched within 30 days are 

subject to a storage fee of $0.05 cents per plant per day; plant materials not claimed within 45 days of notification of 

readiness are forfeited.

IV. Conclusion

This agreement, executed in duplicate, sets forth the entire contract between the parties and may be canceled, 

modified, or amended only by a written instrument executed by each of the parties thereto.

This agreement shall be construed as a contract under the laws of Hawai‘i (name of State or territory and 

country).

Witness the hands and seals of the parties hereto, each duly authorized, the day and year first written above.

______________________________________________

Maria Planter Date

Upland Ranch and Forest

______________________________________________

Gloria Green Date

Green’s Tree Nursery

Figure 18.10—Clients often enjoy staying informed about their 

crop’s progress. Sending e-mail updates with photos or inviting 

clients to visit their crop helps keep clients in the loop. Photo by 

Douglass F. Jacobs.

Sample Client Update

Re: Update on your order for 500 koa 

(Acacia koa)

Date: November 10

Dear Maria,

I hope this note finds you well. I’m writing to 

let you know your 500 koa seedlings are doing 

great—we moved them into the final hardening stage of production last week, so they are 

getting toughened up for outplanting. We are 

right on schedule. Let’s arrange a pickup date 

for some time between December 10 and 

December 15. I think you said you have a van to 

transport them? If not, let me know, and we will 

find a safe transport solution. Please write or 

call back to confirm that you got this message, 

when you get a chance. I look forward to hearing 

from you.

—Gloria, Manager, Green’s Tree Nursery

Continued from previous page
Staff should remember that tremendous effort and 

expense go into planning a project and preparing land for 

outplanting. Acquiring plants is a central part of this process but may be a small percentage of the total project cost. 

Nursery staff should do everything in their power to meet set 

schedules. If any problems or delays are anticipated with the 

crop, clients must be updated immediately so they can modify 

their plans accordingly.

Educating Clients About Shipping, 

Handling, and Outplanting

As described in Chapter 16, Harvesting and Shipping, 

plants are vulnerable to damaging stresses during the transition from the nursery to the outplanting site. Some nurseries take charge of shipping and handling, others leave it 

to the client to pick up and transport their plants. In most 

cases, clients will do their own outplanting. Many clients 

will be knowledgeable about plants and treat them properly 

to ensure success after leaving the nursery. Other clients, 

however, may compromise their plants’ quality by stressing 

them during transport, delaying planting by weeks without 

watering, or planting improperly. Everyone loses when these 

problems occur: the client is unhappy (and may blame the 

nursery), the nursery’s hard work to cultivate the plants is 

wasted, the plants do not survive or thrive, and the environment does not benefit from the presence of the plants. These 

problems can be avoided by educating clients in advance so 

they know how to properly transport, store, and outplant 

their new plants (figure 18.11).

Well before the delivery date for plants, provide clients 

with information to help them properly plan for the best 

shipping, handling, storage, and outplanting practices after 

they receive their plants. For example, you could send them 

copies of Chapter 16, Harvesting and Shipping, and Chapter 17, Outplanting, from this manual, or you could create 

a checklist of “do’s” and “don’ts” based on those chapters 

and your own experiences. These educational materials will 

also help clients communicate best practices to everyone 

who will work with the plants after they leave the nursery.

Following Up With Clients

The success of the nursery, the clients, and the plants 

in the field are interrelated. After an order is complete, 

you need to have a system in place whereby clients can 

provide feedback. Sometimes nursery managers are 

reluctant to follow up because they are concerned that 

they will be blamed for losses or problems. With a good 

contract in place (as described previously), however, the 

limits of the nursery’s responsibility are clearly defined.

Often, success of reforestation and restoration projects 

is monitored after outplanting by tracking survival or other 

important characteristics, as described in Chapter 17, Outplanting. Clients may also keep track of specific problems 

affecting plant survival and growth in the field (pests, 

grazing animals, drought, and so on). This information, 

whether formal or observational, should be shared with 

the nursery.

Working With People 347

Figure 18.11—Educate clients 

in advance so they know how to 

properly transport, store, and 

outplant their new plants. Photo 

by Brian F. Daley.
This field information is important for learning and 

improving nursery practices and is a key part of customer 

relations. Although some failures and problems are inevitable, you will also get an opportunity to see successes, 

which will add to your satisfaction with your work. Ideally the nursery contact person can visit the outplanting 

site and check on the progress of the plants over time. 

Observations can then be used to improve target plant 

specifications for that outplanting environment.

Working With the Community

Community education and outreach are important 

activities for many tropical plant nurseries. The perpetuation of native and culturally important plants is often 

a part of the nursery’s mission, but the best efforts are 

lost if the community is not ready, willing, or able to use 

plants the nursery will produce. Hosting tours, workshops, or field days, writing educational materials, articles, or blogs, and attending local fairs, farmer’s markets, 

or trade shows can all help educate the community about 

the nursery’s mission and availability of plant materials 

being produced. People who might one day plant species 

from your nursery would benefit from learning interesting details about the plants, such as their interactions 

with pollinators or their traditional medicinal properties, 

as well as some of the more practical aspects of working 

with the plants, such as how to plant a tree or the importance of locally adapted seed sources (figure 18.12).

348 Tropical Nursery Manual

Figure 18.12—Hosting workshops or field days helps educate 

the community about the nursery’s mission and plant materials 

being produced, such as this group of university students learning 

about reforesting Hawaiian dryland native species. Photo by Yvonne 

Yarber Carter.

Figure 18.13—Discussing the role of native plants in the local 

forest ecology is an engaging way to involve youth groups. Shown 

are members of the St. Croix, Virgin Islands 4-H club hiking on a 

trail that leads to a forest restoration site where native trees were 

planted. Photo by Brian F. Daley.

Figure 18.14—Sharing knowledge of local plants and their 

uses with future generations is a goal shared by many native 

plant nurseries. Photo by Thomas D. Landis.
Many tropical plant nurseries work with school 

groups and environmental or cultural education activities, such as ecohikes and forest restoration site visits 

(figure 18.13). Connecting with living, growing plants 

can be a wonderful activity for both youth and adults 

and a meaningful way to pass traditional and scientific 

knowledge about plant species, especially their uses and 

benefits, to younger generations (figure 18.14). These 

activities can also increase the community’s desire to 

plant and care for plants from the nursery.

References

Landis, T. D. 2011. The Target Plant Concept—a history and 

brief overview. In: Riley, L.E., Haase, D.L. and Pinto, J.R.tech 

coords. National proceedings: forest and conservation nursery 

associations—2010. Proceedings RMRS-P-65. Fort Collins, CO: 

US Department of Agriculture, Forest Service, Rocky Mountain 

Research Station, Proceedings: 61-66.

Westrum, R. 1994. An organizational perspective: designing 

recovery teams from the inside out. In: Clark, T.W.; Reading, 

R.P.; Clark, A.L., eds. Endangered species recovery: finding the 

lessons, improving the process. Washington, DC: Island Press: 

327–349.

Working With People 349
350
351

Tropical nursery management includes all aspects of growing plants through all 

their growth phases as described in Chapter 4, Crop Planning: Propagation Protocols, 

Schedules, and Records. Management involves an understanding of practical, scientific, technical, interpersonal, and economic aspects of the nursery. Nursery management includes ordering materials and supplies, maintaining facilities, scheduling 

activities, keeping horticultural and financial records, and much more (figure 19.1). 

This chapter outlines some essential elements of the day-to-day and year-to-year 

aspects of managing the nursery.

Nursery Management

Kim M. Wilkinson and Thomas D. Landis 19

Facing Page: Planning space and facilities is an important part of nursery management. Pictured here, staff at the main 

nursery on Palau hook up new irrigation lines. Photo by George Hernández.

Figure 19.1—Nursery management includes not only growing plants, but keeping horticultural and financial records, managing supplies, planning activities (as shown here in 

American Samoa), and much more. Keeping good records is essential to keeping production on track. Photo by Ronald Overton.
Identifying Nursery Tasks

A checklist can help provide an overview of the interrelated tasks involved in managing a nursery. The example checklist in the textbox on the following page can be 

modified and customized to describe your nursery’s daily, 

weekly, monthly, and seasonal tasks.

Only a few required tasks must happen each day; these 

tasks are the essential activities that keep the crops alive 

and healthy and the nursery functioning on a daily basis. 

These tasks include watering, keeping daily records, and 

monitoring crops as they go through the establishment, 

rapid growth, and hardening phases. Other tasks need to be 

done less frequently but are as important. Good planning 

and oversight will ensure that all nursery tasks are prioritized and scheduled accordingly.

Planning and Scheduling

Schedule an overview and planning session on a 

weekly basis to assess immediate needs, periodic tasks, 

and long-term goals. This assessment provides an opportunity to prioritize tasks for the coming week and month 

(figure 19.2). The needs of the plants, environmental 

conditions, and many other factors require flexibility 

and responsiveness in management style. Crops usually 

do not respond well to a rigid schedule and may perform 

differently in different years, which is why weekly assessment and planning is so important. Attempts to make 

rigid schedules (such as “weed every Tuesday”) are often 

far less effective than regular planning to tailor tasks to 

the observed needs and conditions of the crop.

The observation skills of the nursery manager and 

staff are the greatest assets to effective planning. Taking 

time on a weekly basis to review the daily log, plant development record, and other observations will help with prioritizing the work to be done. Observations in the nursery 

can answer many questions: What growth phase is the 

crop in: establishment, rapid growth, or hardening? Is it 

on schedule? What needs to be done next: transplanting, 

moving to a new structure, altering fertilization rates? 

Are we observing anything that might indicate a potential 

problem, such as the presence of a potential pest? When 

do we next update clients on the crop’s progress? Is it time 

to sow a new crop? These observations aid the nursery 

manager to plan and schedule important activities and to 

assign roles, tasks, and deadlines to the appropriate staff 

members.

Some proactive planning should also occur, focusing 

beyond what is most urgent. Keeping the nursery’s vision 

and objectives as a focus during meetings can help maintain a broader view for nursery activities. Planning should 

include activities that are important to the nursery’s 

larger mission beyond the day-to-day details such as public relations or educational activities (figure 19.3). Updating plant protocols and working to improve plant quality 

with some simple experiments are also valuable activities 

that should be included in the nursery’s schedule.

352 Tropical Nursery Manual

Figure 19.2—Organized management oversees the growth of 

the plants and the growth of the nursery itself. A nursery manager 

discusses constructing seedling benches in an area with morning 

sunlight in the U.S. Virgin Islands. Photo by Brian F. Daley.

Figure 19.3—Periodic educational activities help the nursery 

staff to expand their knowledge and skills towards achieving the 

nursery’s mission. Here, nursery professionals meet for a workshop in American Samoa. Photo courtesy of Diane L. Haase.
Nursery Management 353

Planning and Scheduling (weekly, monthly, yearly)

• Make a list of what needs to be done based 

on daily observations, daily logs, and crop 

development records.

• Establish propagation protocols.

• Create and update crop-growing schedules 

and facilities schedules.

• Prioritize and delegate tasks.

• Follow up to ensure tasks were done.

• Refine nursery vision and objectives annually; 

anticipate new crops to grow, changes in 

production, infrastructure improvements or 

expansion, and other planning for the future.

Routine Tasks (daily)

• Irrigate

• Crop culturing (for example, weed or pest 

control, fertilizing).

• Monitor and observe the crops.

Recordkeeping 

• Record observations and actions in daily 

journal or log (daily).

• Make notes in the plant development records 

for each crop (daily or weekly).

• Update and revise plant protocols 

(at end of each crop).

• Conduct crop inventory assessment (ongoing).

Crop-Phase Production Tasks 

• Establishment tasks (for example, making 

growing media, sowing seeds, inoculating 

with microsymbionts).

• Rapid growth phase tasks (for example, 

fertilizing, monitoring).

• Hardening phase tasks (for example, changing 

fertilization and light regimes).

• Update clients about crop development.

• Harvesting, packing and shipping tasks.

Seasonal Cleanup 

• Purge or transplant holdover stock.

• Clean floors, tables, tools, equipment, and so on.

• Clean and sterilize containers.

• Check and repair equipment, tools, and 

infrastructure such as irrigation lines.

Financial Management

• Determine expenses, including labor, time 

and supplies needed to produce crops, and 

overhead costs (for example, utilities).

• Determine estimated income.

• Create and manage an annual budget based on anticipated income and expenses.

• Administer contracts.

• Inventory supplies for production and maintenance 

(for example, growing media, fertilizers, containers and 

trays, irrigation parts) and order as needed.

• Estimate future costs and income and adjust 

budget accordingly.

Problem Solving and Troubleshooting

• Identify and analyze problems as they arise.

• Know who to call for help (for example, another grower, 

a soil scientist, a pest expert, an irrigation specialist) 

and contact them as needed.

• Develop and test hypotheses to solve the problem.

• Implement a solution.

Cultivating Good Relationships with Staff, Clients, 

and the Community

• Provide staff education and training.

• Connect staff with nursery vision and objectives.

• Give and receive feedback and input (observations and 

improvement suggestions).

• Plan meetings, safety awareness, and so on.

• Develop target plant specifications with clients.

• Educate clients about key issues for handling, outplanting, and care.

• Visit outplanting sites and clients to check up on 

survival and growth; follow up with clients to discuss 

field performance of plants, revisions of target plant 

criteria, and future needs.

• Offer public education and outreach.

Learning and Sharing

• Attend training events and conferences.

• Learn from other nurseries; host and attend field days 

and visits.

• Read published literature (for example, Native Plants 

Journal).

• Explore ways to improve crop production and plant 

quality.

Nursey Management Checklist
354 Tropical Nursery Manual

Daily Tasks and Observations

Daily activities include the essentials of keeping the 

plants alive and healthy such as irrigating, weeding, 

monitoring, fertilizing, and pest management (figure 

19.4). Daily observation of the crop is essential for good 

nursery management and an important way to determine their needs.

The manager or a designated “crop monitor” needs to 

observe the crop every day (figure 19.5). This task can be 

done once daily as a formal practice separate from other 

tasks. Measurements occasionally will be taken to evaluate the crops progress or to quantify environmental conditions. The person monitoring the crop needs to understand 

what “normal” is for that crop and for the nursery environment and be highly sensitive to any deviations from that 

norm (figure 19.6).

Experience and daily observations can identify potential problems long before they become emergencies. 

Observations may include the following (Landis 1984)—

• Appearance—Inspect the crop visually to see if plant 

size and shoot-to-root ratio are proper for the stage of 

growth. Look for signs of nutrition or disease problems 

on the roots and foliage. Inspect closely for insect pests. 

Examine roots to see if beneficial microsymbionts are 

visible.

• Smells—Some problems, such as gray mold, may 

be discernible to the experienced grower by its 

odor. Overheating motors, broken fans, composting 

troubles, and other potential problems can also be 

detected by the sense of smell.

• Noises—Listen for unusual sounds in the nursery 

such as an engine running unsteadily or water running 

when it should not be.

• Feel—Pay attention to the temperature and humidity 

in the nursery. Feel the growing medium to determine 

if it is at the proper moisture level.

Figure 19.4—Daily activities include the essentials of keeping the plants alive and healthy, including watering (A), weeding (B), and 

timely transplanting (C). Photo A in Yap by Tara Luna, photo B in Tanzania by Ronald Overton, and photo C in U.S. Virgin Islands by 

Brian F. Daley.

B

C

A
Nursery Management 355

Figure 19.5—A manager or a designated “crop monitor” 

should observe the crop every day. Photo at Volcanoes National 

Park, on the Big Island of Hawai‘i, by Brian F. Daley.

Figure 19.6—Making regular observations of the crop and 

staying alert and aware are key to avoiding problems. Photo by 

Tara Luna.

Although one designated crop monitor is responsible for 

this task, all staff need to understand that observation and 

being alert and aware are key to precluding problems. The 

manager should welcome and encourage staff to share their 

observations and contribute to the daily log; this practice 

builds observation skills and greater crop awareness.

Recordkeeping

As described in Chapter 4, Crop Planning: Propagation 

Protocols, Schedules, and Records, keeping good records 

is essential to keeping production on track and precluding serious problems (figure 19.7). Nursery recordkeeping 

includes the following—

• A Daily Log—can be as simple as writing the day’s 

date and jotting down some notes in a notebook about 

observations and activities at the end of each day. Make 

a habit of entering something in the log each day, even 

if the observations seem unimportant at the time.

• Plant Development Records—for each crop should 

be kept in an easily accessible place and a few notes 

should be jotted down as changes occur with the crop.

• Propagation Protocols—show how to produce 

each kind of crop successfully in your nursery. The 

protocols provide guidance for each new crop in 

developing the production plan and listing needed 

facilities and supplies and should be regularly 

updated and revised.

• Inventory Assessment—should include all plants 

in the nursery by bench or structure number, current 

developmental stage of each crop, and details of 

delivery (target requirements, outplanting site, name 

of client, seed source, and anticipated delivery date).

As the nursery grows in size and complexity, entering 

records into a computer will make the information easier 

to track. Every day creates small amounts of vital information that will contribute immeasurably to improving nursery 

management and productivity over time (figures 19.8, 19.9). 

Figure 19.7—The daily log, the plant development record, and the 

propagation protocol are the foundation of nursery recordkeeping 

and are used to support plant production and inform nursery management. Illustration adapted from Dumroese and others (2008) by 

Jim Marin.
356 Tropical Nursery Manual

This information is invaluable for many aspects of nursery management, including the following—

• Budgeting funds.

• Estimating schedules to produce future crops.

• Determining what labor-saving equipment might give 

the most benefit for the cost.

• Analyzing nursery expenses.

• Improving profits or production.

• Replicating successful crops.

Crop Planning and Production 

Tasks

The details of planning crops are discussed in Chapter 

4, Crop Planning: Propagation Protocols, Schedules, and 

Records. Management needs during the crop production 

include—

• Understanding the three growth phases crops go 

through (establishment, rapid growth, and hardening) 

and the distinct requirements for each phase.

• Making growing schedules for crop production 

from seed procurement through outplanting and 

detailing changes as the growing cycle progresses 

(figure 19.10).

• Listing space, labor, equipment, and supplies required 

to support the crop during the three stages of growth.

The work to produce a crop consists of managing the 

plants through each phase of development so that plants 

receive what they need and are as strong and healthy as 

possible for outplanting. After a schedule has been made 

showing what plants need to be sown and by when, the 

tasks of preparing growing media, filling containers, and 

sowing can be scheduled. In the establishment phase, 

plants begin to germinate, and thinning, transplanting, and 

inoculation with beneficial microorganisms will take place. 

As the plants move from the germinant phase to the rapid 

growth phase and later the hardening phase, their needs 

will change.

For some nurseries, plants will be physically moved from 

a germination or rooting area to a more open environment 

(figure 19.11). For other nurseries, climate control (such as 

removing shadecloth) might have the same effect. FertilizaFigure 19.8—Recordkeeping tools should be easily accessible at 

the manager’s workstation; make a habit of entering something in it 

each day. Photo by Kim M. Wilkinson.

Figure 19.9—The timing of nursery tasks, such as transplanting, 

should be recorded in the daily log and the plant development log. 

These records will help you keep the current crop on track and will 

help you plan, budget, and schedule for future crops. Photo by Kim 

M. Wilkinson.

Figure 19.10—The work to produce a crop consists of managing 

activities and resources through each phase of crop development. 

This whiteboard at the Guam Division of Forestry and Soil 

Resources nursery lists crops being grown for the Cetti Bay 

restoration project including how many plants are needed 

(species and container types) and the current status of the crops. 

Photo by Ronald Overton.
Nursery Management 357

tion and watering regimes are changed for each of the three 

phases. When the crop is ready, it will be harvested and 

shipped, as described in Chapter 16, Harvesting and Shipping.

To keep up with production, planning for space and 

facilities is important. The manager also needs to have the 

necessary materials on hand when they are needed (figure 

19.12). These materials include supplies for production 

such as seeds, growing media, containers, and so forth. For 

nurseries in remote areas, obtaining supplies may require 

ordering months ahead of the time when they are needed. 

In these cases, extras of essential items need to be kept on 

hand. If spare parts are used for repairs, they need to be 

replaced right away.

Seasonal Cleanup and 

Maintenance

Time is usually available between crops or at the end 

of each season for some “deep cleaning” and maintenance. 

Cleanliness is essential to avoid disease problems and to 

maintain a professional, appealing image for the nursery. 

A clean environment builds customer confidence and staff 

morale. Perform the following tasks every 2 to 6 months:

• Dispose of any holdover stock.

• Clean and hose down the floors and tables 

(applying dilute bleach or other cleanser if no 

plants are present).

• Clean and sterilize containers (see Chapter 7, 

Containers).

• Flush out irrigation system and run a cup test 

(see Chapter 11, Water Quality and Irrigation).

• Conduct equipment checks and make any repairs.

• Replace roof plastic, if necessary.

Financial Management

Good financial management is necessary for the nursery is to thrive in the long term. The daily journal and other 

records should include the amount of labor spent on various 

activities, money spent for materials, and overhead costs such 

as rent and utilities. These costs can be used to estimate the 

cost of each crop, thereby enabling the manager to accurately 

budget time and funds. This information also is essential for 

determining what the nursery must charge for various plant 

materials sold. Learning more about basic business planning 

through publications and local learning opportunities can be 

very helpful for making wise financial choices.

Figure 19.12—Management involves planning the timing of essential production tasks and ensuring that all the necessary supplies 

are available. Shown here: checking a seed supply in Hawai‘i (A), filling bags with growing medium made from composted coffee 

cherries in East Timor (B), and assembling a drip irrigation system on Yap (C). Photo A by Kim M. Wilkinson, photo B by J.B. Friday, 

and photo C by George Hernández.

Figure 19.11—Plants are moved from one structure to another 

as they go through their three phases of growth. Good planning and 

management maintains open paths and an easy flow of work between structures. Photo by Tara Luna.

A B C
358 Tropical Nursery Manual

In keeping financial records, be sure to note the following factors:

• Size of stock and growing space needed to produce it.

• Time to grow.

• Labor (in person-hours) required through all phases.

• Materials required and their cost (for example, 

seeds, containers, growing media).

• Need for custom culture (for example, special 

containers, extra labor).

• Overhead costs (for example, utilities).

• Cost inflation over time.

• Typical losses (percentage of crop discarded).

• Price charged when the crop left the nursery.

Problem Solving

Good management, staff training, monitoring, and 

planning will generally preclude emergency situations in 

the nursery. Even the best manager, however, cannot avoid 

problems entirely. Some problems, such as difficulties 

with the irrigation system, appear suddenly and must be 

handled instantly. Others require a longer term approach. 

With experience, troubleshooting problems may become 

easier. Do not be reluctant to reach out to colleagues, other 

nursery managers, or other professionals. Everyone has 

problems once in a while, and we can all help each other 

learn more about plants as we share our experiences. 

The problem should next be classified into some sort 

of order, separating problems into three types based on 

the amount of information available (Landis 1984, Van 

Gundy 1980):

• Type I problems are well structured. These are the 

routine problems that occur daily. Their main characteristic is that all the information needed to solve 

them is already available. Problems in this category 

have probably occurred before and can usually be 

solved by standard procedures. Expertise for solving 

these problems can normally be found at the nursery, 

so outside help is not required.

• Type II problems are semistructured. This is an 

intermediate category—some information about the 

problem is available, but some degree of uncertainty 

also exists. These problems may have occurred before 

but something about them makes them different. 

Existing techniques must be adapted to solve this type 

of problem, and some expert help may be needed. The 

final solution is probably a combination of standard 

and newly developed methods.

• Type III problems are poorly structured. Their distinguishing characteristic is that little or no information 

is available about them. These are the problems never 

encountered before; therefore, expert help should 

be sought and the information needed to solve these 

problems generated through the problem-solving 

process. Solutions to poorly structured problems 

usually have to be custom made and require creative 

problem-solving techniques.

Type 1 problems are usually solved with standard operating procedures, whereas Types II and III require more 

creative steps.

Figure 19.13—A problem-solving matrix. Illustration adapted 

from Landis (1984) by Jim Marin.
Nursery Management 359

This five-step systematic approach can be helpful when 

approaching long-term challenges (figure 19.13; Landis 

1984):

• Identify the Problem—Is it really a problem? What 

seems to be wrong?

• Analyze the Problem—What happened exactly? 

When did it start?

• Generate Ideas—Identify potential sources of the 

problem. Consult literature, other nurseries, staff 

members, or outside sources of help such as extension 

agents or specialists.

• Develop and Test Hypotheses—At some point, a 

conclusion about the source of the problem must be 

decided and acted upon.

• Implement a Solution—Decide on a way to solve the 

problem. Observe the results. If the problem is not 

solved, start again with step 2.

References

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p. 

Landis, T.D. 1984. Problem solving in forest-tree nursery with 

emphasis on site problems. In: Duryea, M.S.; Landis, T.D., eds. 

Forest nursery manual production of bareroot seedlings. The 

Hague, The Netherlands: Martinus Nijhoff/Dr. W. Junk Publishers: 307–314. 

Van Gundy, A. B. 1981. Techniques of structured problem solving. Van Nostrand Reinhold, New York. 301 p.
360
361

Working with plants is a process of discovery. Being curious and aware, paying 

close attention, and staying open and adaptive are important practices. Books and 

people can help us learn about plants in the nursery, but the very best teachers are the 

plants themselves. “Research” is simply tracking what is happening, asking questions 

about what is causing it to happen, and seeking answers. In other words, research is 

something most growers already do. Astronomer Carl Sagan (1996) said, “Every time 

we exercise self-criticism, every time we test our ideas against the outside world, we 

are doing science.” Scientific research is simply “the testing (systematic, controlled, 

empirical, and critical investigation) of ideas (hypothetical propositions about presumed relations among natural phenomena) generated by intuition” (Dumroese and 

Wenny 2003). If research is done well, the process can yield useful, accurate information. The purpose of this chapter is to provide information about how to design easy 

trials and experiments to discover useful, meaningful ways to improve crop production and quality.

Some people may be lucky enough to have an elder or mentor pressing them to 

explore and discover more. Carrying out research, asking questions, and keeping 

records are ways to self-mentor. Using a systematic approach supports making accurate observations so that discoveries otherwise missed can be made and shared. It is 

widely acknowledged that working with plants is an art as well as a science: observation, senses, emotions, empathy, and intuition play important roles. When a question arises that is important to answer, it is time to consider conducting a trial or 

experiment. Growers often work with plants for which minimal literature or outside 

information is available. Nursery research may be subjective or objective, simple or 

complex. Learning how to effectively carry out experiments to evaluate new plant production techniques is essential to discovering relevant and applicable practices.

Managing a nursery in the tropics is a profession where you will learn as much or 

more from your own direct experience than you will learn from readings, classes, and 

other formal teaching. Some learning comes from experiments; but some comes from 

experience and reflection. A few tips for reflective practice as a way to continually learn 

and adapt are provided at the end of this chapter.

Discovering Ways To 

Improve Nursery Practices and 

Plant Quality

Kim M. Wilkinson and Diane L. Haase

20

Facing Page: A germination experiment. Photo by Brian F. Daley.
The Importance of Trying New 

Things in the Nursery

Often, the tendency is to take the path of least resistance 

and use known or established nursery production techniques. The first propagation technique that was tried and 

that produced an adequate plant may have become the established protocol. The technique, however, may be more costly 

or inefficient than alternate methods, and may not produce 

the best quality plant for the outplanting site. A few modifications could improve production, plant quality, and, ultimately, plant survival and growth after outplanting. Simple 

experiments enable the nursery to try out new techniques, 

ideas, and problem-solving strategies (figure 20.1).

The tasks of keeping up with day-to-day nursery management may feel like more than enough to fill your schedule, and time for experimentation may seem a low priority. 

In truth, however, most growers already engage in investigations on a regular basis. Experimentation happens every 

time a new idea is tested, a question leads to alternative 

strategies, or a problem is analyzed and solved. Taking a 

little care to be systematic and follow a few guidelines will 

dramatically increase the benefits of these activities and 

will provide greater confidence in the conclusions you 

draw from them.

Simple experiments can be carried out simultaneously 

with filling an order for plants. For example, in most cases, 

the nursery can produce plants using an established protocol and also grow some additional plants for research at 

the same time. One variable can be altered for the experimental plants, such as using a different seed treatment, a 

modified growing medium, a new type of container, or a 

different mycorrhizal inoculant. Your results for the experimental group can then be compared against those for the 

established protocol. In this way, each new crop represents 

an opportunity to try something new on a small scale. The 

discoveries can greatly improve production efficiency and 

seedling quality over time. Because of these potential benefits, it is worth putting a little effort into experimenting 

and trying new things.

362 Tropical Nursery Manual

The Value of Experimenting

Today’s best methods for raising native plants came 

from growers like you learning through experimentation, then sharing what they learned. For example, 

Kerin E. Lilleeng-Rosenberger describes her process: 

[in 1989] “I was hired and put in charge of the new 

native Hawaiian plant nursery...I was provided with 

a variety of native seeds, but had neither instructions 

nor reference guides to follow; the majority of these 

plants had never been grown in cultivation before...

For more than ten years, I experimented with different potting mixes, seeds, and cutting treatments, 

along with many other types of plant materials. I 

kept detailed records, which helped identify what 

had contributed to the successful results. Eventually, 

the knowledge I had gathered filled fourteen notebooks.” Her book, Growing Hawai‘i’s Native Plants, 

shares what she learned. 

(Lilleeng-Rosenberger 2005)

Figure 20.1—Simple experiments and trials, such as this fertilization trial, can teach us a lot about how to grow plants. 

Photo by R. Kasten Dumroese.
Conducting simple experiments and trials in the 

nursery can accomplish the following—

• Produce better plants.

• Speed up production.

• Save money, labor, seeds, and other materials.

• Improve outplanting survival and performance.

• Refine target plant criteria for outplanting sites.

• Contribute to greater knowledge of the plants.

Making Observations and 

Keeping Records

A foundation for improving plant health and quality 

is a good understanding of current production practices 

and how plants respond to them. The following three 

basic types of records, as discussed in Chapter 4, Crop 

Planning: Propagation Protocols, Schedules, and Records, 

are the foundation of good nursery recordkeeping (figure 

20.2):

1. A daily log of general conditions and activities.

2. Development records for each crop that are filled out 

as the plants develop.

3. Propagation protocols that describe from start to finish how the plants are currently grown.

Additional records may also be enjoyable and valuable, such as an informal, personal journal that documents 

reflections in growing plants or a photo diary.

These records prevent the nursery from wasting time 

repeating strategies that do not work while providing a 

plan to help duplicate successful crops. This information 

also establishes how crops normally perform and can be 

used to recognize problems or gauge the effect of alternative production methods. Perhaps most important, keeping data and cultural records in a written format creates 

information that can be passed on to future nursery staff 

or others in the community. Without these records, valuable information (perhaps gleaned from a lifetime of nursery work) may be lost, and the new grower will have to 

start over. Knowing in a measurable way what is “normal” 

helps the nursery experiment with modifications that can 

improve crops and productivity over time.

Discovering Ways To Improve Nursery Practices and Plant Quality 363

Figure 20.2—The nursery’s daily log, plant development record, 

and propagation protocol are the three basic kinds of records that 

form the foundation for learning. Illustration adapted from Dumroese and others (2008) by Jim Marin.

Solving Problems 

Through Collaboration and 

Questioning Assumptions

In our small research nursery, we had poor results 

growing the Virgin Islands native tree Clusia rosea

(in some places known as the “autograph tree”). 

We kept having low germination rates. I shared this 

experience with my friend who is the local nursery 

owner at Cruzan Gardens. He offered to give the 

seeds a try. When he reported a nearly 100-percent 

germination, I knew he found “the trick.” What had 

he done differently? 

The tiny seeds of Clusia rosea are embedded in the 

flesh of a succulent fruit and often are eaten by bats. 

My standard protocol for all tree species at the time 

included removing all fruit and flesh from seeds before 

germination. My friend, however, found this method 

too time consuming and simply spread the seeds, still 

embedded in their fruit pulp, into trays of potting mix, 

as if he were spreading jam on toast! It turns out that 

my method was dessicating the tiny seeds that are not 

normally exposed to air.

Naturally, we revised our protocol to keep the 

seeds in their pulp during germination, and we were 

able to get great germination results, too. I learned 

from this experience to share my challenges with 

other growers and to question my assumptions.

—Brian F. Daley, U.S. Virgin Islands
Step 1: Developing Ideas 

for Nursery Experiments

Ideas for experiments may come from many places, 

including the following—

• A casual observation the nursery wishes to verify.

• A pressing question that seems to recur in the daily 

log or journal.

• An informal trial in which a difference is observed.

• A need to improve something (for example, percentage of germination).

• A need to change practices or materials (for example, 

the rise in cost of a potting media ingredient requiring 

use of substitutes).

• A desire to work with a new species.

• A desire to try out a new technique or idea.

Because staff time is limited, ideas for experiments 

will likely be prioritized in terms of their potential positive effect and importance to production. Nevertheless, it 

is good to keep a list of any potential experiments or trials 

that you or other staff believe would be beneficial and keep 

these ideas on hand to try as time allows. As described 

364 Tropical Nursery Manual

We grow an important Hawaiian endemic tree, the 

‘ōhi‘a tree. This tree is a foundation of many Hawaiian 

native ecosystems. A problem we had is ‘ōhi‘a rust (figure 20.3). ‘Ōhi‘a rust is easily spread by the wind, and 

our nursery in Kamuela, on the north tip of the Island 

of Hawai‘i, is always windy.

The disease would get so bad at our nursery that our 

mortality rate was quite high. So we started observing the 

‘ōhi‘a plants in different locations around the nursery. 

Although not fully scientific in our methods, we noticed 

some things that caused us to rethink the plant locations 

throughout our nursery.

Basically we found that—

• Plants in our greenhouse were very susceptible to 

the rust disease (which may be because the wind 

slows down and “circles” in the greenhouse because 

only one side of the building is open, causing the wind 

to blow the rust spores gently around in a circular 

pattern over all the plants in the greenhouse).

• Plants outside exposed directly to the wind (such as on the windward side of a table or pallet) seemed as 

susceptible as the plants in the greenhouse.

• ‘Ōhi‘a seedlings located outside on the leeward side of larger plants were relatively rust free. The larger 

plants may cause a disruption in the wind pattern, pushing the rust up and away from the smaller plants 

behind them.

After we observed these three things, we adjusted the locations of our ‘ōhi‘a seedlings so they would be protected 

from the wind, essentially hidden from the rust. It did not totally eliminate the problem, but we experience far 

lower mortality rates and fewer diseased plants. We can tolerate the amount of loss now. We have tried this 

approach with limited success with other diseases and problems such as moving plants susceptible to mildew or 

powdery mold to be surrounded by plants that do not seem nearly as susceptible. So far, however, nothing has been 

quite as dramatic or successful as “hiding” our ‘ōhi‘a seedlings from the ‘ōhi‘a rust.

—Jacob Witcraft, Nursery Forester, State Tree Nursery, Kamuela, Hawai‘i

Figure 20.3—‘Ōhi‘a rust, shown here on an ‘ōhi‘a leaf, 

is a disease spread by wind. Observations of wind patterns 

and experiments with plant locations helped staff at the 

State Tree Nursery in Kamuela, Hawai‘i, to minimize losses. 

Photo by J.B. Friday.

Solving Problems Through Observation
earlier, some experiments may be easy and quick to carry 

out and can be done efficiently as part of crop production.

Many topics lend themselves well to experimentation, 

including the following—

• Developing seed treatments and 

germination techniques.

• Testing culturing options for new species or 

seed sources.

• Researching microsymbiont sources or 

application methods.

• Altering watering regimes.

• Trying new container types.

• Trying new blends or materials in 

potting media.

• Changing an aspect of management, 

such as timing for moving crops from one 

phase to another.

Step 2: Gathering Information

After you have an idea of what you want to investigate, 

some background research is in order. Check your existing 

plant development records, plant protocol, and daily log to 

review what has happened and has been tried in the past, 

reflecting on your experiences and observations. Information can also be gathered from nursery staff, specialists, 

extension agents, and associates from other nurseries. Look 

up the subject in journals, books, and electronic sources of 

information. This background research can help narrow the 

focus of the experiment to a question that can be answered.

Step 3: Specifying a Question and 

Creating the Hypothesis

The question must be limited in scope, pertaining to 

only one aspect of production. It would not work to focus 

on multiple variables at the same time. For example, if 

more than one aspect of the seed treatment process (such 

as, season harvested, collection and processing method, 

sowing times and methods, and medium used during the 

germination phase) were modified at the same time, how 

would you know which modification caused a difference?

The hypothesis is the proposed answer to the focused 

question.

Step 4: Designing the Experiment

A good study design to answer a specific question and 

test the hypothesis starts with careful selection of the treatments to be evaluated (the independent variable) and the 

Discovering Ways To Improve Nursery Practices and Plant Quality 365

Example: Step 1—An Idea for an Experiment

A native species consistently has less than a 25-percent germination after sowing. In addition, germination 

is sporadic, taking place over 4 weeks. These results 

occur despite good seed sterilization and handling 

practices. A seed test sent to a laboratory indicated 

that a higher percentage of the seeds are viable. The 

nursery wishes to increase the percentage germination. The first step is to look at the options for experimentation. What could be contributing to the low 

germination for this species in your nursery? And 

what could increase germination?

Example: Step 2—Gathering Information

An Internet search, along with a review of this 

manual’s Chapter 8, Collecting, Processing, and Storing Seeds, indicates that when viable seeds fail to 

germinate, the seeds may be dormant and, therefore, 

may require a treatment to break dormancy. A bit of 

reading into the natural history of this species reveals 

that its seeds are often passed through the digestive 

system of birds, which can be a form of scarification, 

breaking natural dormancy. According to publications 

and other nursery growers consulted, several closely 

related species have a hard seedcoat that is usually 

scarified before planting. A search on the Native Plant 

Network (http://www.nativeplantnetwork.org/) may 

not find your exact species, but it shows that several 

growers have had success with a few related species 

when using mechanical scarification.

Example: Step 3—Specifying the Question 

and Creating the Hypothesis

From your information gathering, it appears that 

lack of scarification may be causing the poor germination of your species. The question that the experiment 

will address is now formulated: How does scarifying 

the seeds of this species affect its germination? The 

hypothesis might be, “Mechanical scarification of the 

seeds by hand nicking the seedcoat with a small nail 

clipper will result in improved germination.” If time 

allows, it may be desirable to pose more than one 

treatment to answer the same question. Perhaps a second treatment to be tested separately for this example 

would be, “Hot water scarification for 20 seconds will 

result in improved germination.”
response to be measured (the dependent variables). 

From there, the experimental design can be decided. 

The “Three R’s” of designing a good experiment are Replication, Randomization, and Representation and are 

described in the following sections. Even the simplest 

study design needs to include these important elements 

to minimize the risk of generating incorrect or meaningless data. The components of the study including the 

question, hypothesis, plant material, treatments, and 

experimental design should all be written into a study 

plan. The study plan needs to read like a recipe so that 

you know what, why, when, where, and how to conduct 

your study.

Independent Variable 

(Treatments)

The independent variable is the one factor purposely 

changed for the sake of the experiment. It is the factor 

that is expected to create a response. A factor could be 

seed source, seed treatment method, growing medium, 

irrigation frequency, fertilizer rate, and so on. But, only 

one needs to be used as the independent variable in a 

simple nursery trial; all other factors must stay the same. 

This approach enables you to isolate whether or not 

changes in that factor result in a desired response. The 

independent variable can be modified one or more ways 

(treatments) and compared with an unmodified treatment (the control).

The Control

An essential aspect of experimenting is to have a control 

treatment. For experiments in plant production, the control treatment is simply the way the nursery usually does 

things according to established protocols. The performance 

of control plants grown in the usual way will be compared 

with the performance of plants grown in the modified way. 

All plants in the experiment need to be started at the same 

time and kept in the same areas of the nursery to eliminate 

further variation among them, thereby isolating effects of 

the independent variable (figure 20.4).

Dependent Variables

The dependent variables are the variables being 

observed or measured. These are the variables that are 

hypothesized to be influenced by the independent variable. Although many variables can be affected by specific 

treatments, you will want to decide ahead of time which 

dependent variables are most important to measure. For 

instance, fertilizer rate can change foliar nutrients, growth, 

survival, media chemistry, and so on. However, you will 

want to focus on the variable(s) that address your experimental question and hypothesis.

366 Tropical Nursery Manual

Figure 20.4—Seed scarification and stratification requirements are often discovered through experimentation and trials. 

Careful labeling of the control and each treatment is essential. 

Photo by Tara Luna.

Figure 20.5—Replications need not be large scale but should 

consist of at least 30 plants per treatment (including the control). Photo by Brian F. Daley.
Replication

Research is largely about isolating the independent 

variable and eliminating the possibility that any other factors could be contributing to observed differences in the 

dependent variables. Replication is an essential aspect of 

this process (figure 20.5). Without replication, you cannot 

confidently conclude that your findings are repeatable. If 

too few plants are used in the experiment, any differences 

observed among treatments may simply be a coincidence. 

For this reason, it is valuable to have as many plants as is 

reasonable included in the experiment.

Variability in the nursery environment can profoundly 

affect experiments. For example, two benches may differ in 

the amount of irrigation or light received. Because of environmental variability, it is best to place plants from all treatments 

(including the controls) right next to each other, on the same 

bench or even in the same tray. (Of course, this placement is 

not possible if the objective of your experiment is to compare 

different growing environments; in that case, the plants will 

have to be located separately.) Every effort needs to be made 

to keep all conditions the same except the one being studied. 

Otherwise, you might find differences among treatments that 

are not actually caused by the treatment.

Replications can be individual plants or can be a group 

(block) of several plants (for example, a tray of seedlings). 

When blocking, place each of the replications in different parts of the nursery if possible. That way, if the same 

relative treatment differences are observed within each 

of the blocks, in spite of variations around the nursery, 

you can have greater confidence that the results are not a 

fluke. Each block needs to consist of a similar number of 

plants from every treatment (including the control). For 

example, a block might consist of 10 control plants, 10 

treatment-1 plants, and 10 treatment-2 plants for a total 

of 30 plants in each block.

A few tips for having good replication include the 

following—

• Have at least 30 to 40 plants total per treatment 

(including the control)

• When blocking, use a minimum of 4 blocks and include 

at least 10 plants from each treatment in each block.

• For each replication, situate all treatments (including 

the control) adjacent to each other to reduce variation 

in microclimate.

• If possible, place replications in three or four different 

locations within the nursery.

• Because each growing season is different, the experiment may be repeated one or two times on subsequent batches of plants to confirm results.

Representation

Another important aspect of experimental design is to 

make sure the plant materials and treatments used in the 

experiment are representative of the same plant materials 

and treatments to which you plan to apply your results. For 

example, if you want to learn about the effect of fertilizer on 

seedlings grown in a particular sized container, then you do 

not want to do your study on seedlings grown in smaller or 

larger containers.

Randomization

It is important to randomly select the plants to be 

included in the study from the pool of representative 

plants (or seeds). It is also important to randomly assign 

treatments to each replication (plant or group of plants as 

described previously). Randomization prevents any systematic or personal bias from being introduced into the 

study and skewing the results; for example, using only 

the largest seedlings for one of the treatments would lead 

to the incorrect conclusion that that treatment made the 

seedlings grow more. Many ways exist to randomize. For 

example, you can roll a die or draw from a deck of cards 

to determine which numbered tray of seedlings to use in a 

study or you can draw slips of paper out of a bag with treatment names: control, treatment 1, and so on.

Discovering Ways To Improve Nursery Practices and Plant Quality 367

Example: Step 4—Designing the Experiment

From your research, questions, and hypothesis, you 

have decided to look at scarification as your independent treatment variable. The study design will include 

three treatments: (1) hand nicking the seedcoat with a 

small nail clipper, (2) immersion in hot water (200 °F 

[93 °C]) for 20 sec, and (3) a nonscarified control (the 

current method used by the nursery). The dependent 

variables to be measured will be percent germination 

and rate of germination. You decide to have four replications; each replication will consist of 25 seeds from 

each of the three treatments for a total of 100 seeds 

in each treatment and 300 seeds total in the study. 

From the total supply of seeds, 25 seeds will be randomly selected and then randomly assigned to a block 

and treatment. For each replication, all seeds will be 

treated and sown at the same time into a single seed 

tray. The four seed trays (replications) will be placed 

throughout the nursery’s germination area to see if 

results are similar among treatments.
Step 5: Conducting the 

Experiment

If you feel unsure about the validity of the proposed elements of an experiment, find an ally in the local university or agricultural extension system to discuss your plan 

briefly. This investment of time is wise and ensures that the 

research successfully addresses the question posed.

After the hypothesis is posed and the treatments and 

experimental design determined, it is time to plan when 

and how to carry out the experiment. Unless the problem 

addressed is urgent (that is, interferes with production), it 

may be most economical to wait until you have an order for 

the species you wish to investigate. A group of plants from 

those grown according to the usual protocol will be designated as the control treatment. Extra plants can be planted 

at the same time from the same seed source and on the same 

day, with only the independent variable manipulated.

A few tips for starting and carrying out the experiment 

include the following—

• Have one person in charge of setting up the experiment, making observations, and collecting all data. 

Having one person in charge helps eliminate unnecessary variations in the data.

• If special materials are needed for the experiment (for 

example, a different microbial inoculant, a new seed 

source, a special growing medium ingredient), be sure 

to have them on hand before the experiment starts.

• Mark treatments (including the control) clearly with 

durable, easy-to-read labels. Nothing is worse than 

discovering a group of plants performing outstandingly but with no record of what was done differently.

• Do not count on experimental treatments to produce 

marketable plants. Use established protocols to meet 

client requirements. Plants devoted to research need to 

be above the count required for the order. If the experimental subjects turn out to be of high quality and saleable, that will be a side benefit. If growing on contract, 

the client may be interested in accepting research plants 

and to continue the trial in the field. Agreements need to 

be clarified in advance regarding experimental plants.

• Take careful notes and keep a journal documenting every step of the experiment as it is carried out. 

Changes may occur rapidly and go unnoticed if care 

is not taken to record them. Sometimes the independent variable will affect one brief but important stage 

of plant development. Keep data organized, ideally 

entered into a computer spreadsheet soon after taking 

measurements and observations.

• Be prepared to carry out the experiment more than 

once.

Step 6: Making Observations and 

Collecting Data

When gathering data, keep the process as simple and 

straightforward as possible, and reduce risks of nonappli368 Tropical Nursery Manual

Example: Step 6—Observations and 

Data Collection

Each of the three seed treatments in the four seed 

trays (replications) in the experiment is monitored 

daily for 4 weeks and the number of germinants 

recorded. (If the emergents are to be transplanted into 

larger containers before the end of the experiment, 

it is critical to ensure the counts are accurate before 

transplanting. Germinants from each treatment are 

transplanted in separate trays and carefully marked, 

even in their new containers.) The percent and rate of 

germination are calculated from the collected data.

Example: Step 5—Conducting the Experiment

The experiment on seed scarification will be carried 

out simultaneously with growing an order of plants 

for that species. Do not expect to be able to use any of 

the seeds from the experiment (the new method may 

increase, decrease, or have no effect on germination 

percentages). So, the procedures to produce the correct number of seedlings to meet the order need to be 

carried out as described in your usual protocol for that 

species. If an order for 100 plants of the species with 

the usual expected 25-percent germination is received, 

according to the established protocol, the nursery will 

need to sow about 450 seeds to compensate for the low 

germination and other losses. A portion of those seeds 

(100) are designated to be part of the experiment as the 

control. At the same time, 100 additional seeds from 

the same seedlot would be scarified using mechanical scarification and 100 seeds would be subjected to 

the hot water treatment according to the experimental design. The four experimental seed trays (replications) and the group of 25 seeds of each treatment 

within each tray are clearly marked. All seeds will be 

treated identically otherwise. Because germination 

usually takes place sporadically over a 4-week period 

to achieve a 25-percent germination, the experiment 

will run 4 weeks.
cable or meaningless results. For example, the wet weight 

of live plants will vary considerably depending on irrigation and time of day; therefore, weighing live plants does 

not usually generate meaningful data for experiments. For 

small experiments, have only one person take measurements and gather hard data (such as plant height or stem 

diameter) to reduce variations in the way data are collected. For larger experiments, however, it may be necessary for several people to collect data. If several people 

will be collecting data, make sure each person is trained 

to measure using the same procedures. In these cases, it 

is a good idea for the different people to take data on all 

treatments, including the control, to cancel out bias in 

data collection (instead of having one person collect all 

the data for the control and a different person collecting 

the data for the experimental plants).

Data collection for the experiment needs to focus on 

the dependent variable for that experiment. Other data and 

observations, if available, however, may be collected as well 

if time allows. Even if they are not quantified, observations 

about the appearance and vitality of plants can be especially 

useful for many experiments. Taking several photos of the 

experiment is another way to document your findings. 

Remember, by doing this type of work, you are gaining 

expertise; your observations are meaningful.

The best timing for data collection varies depending on 

what is being studied. Although any period of rapid change 

for the crop can be a useful time to gather data, in general, 

the most meaningful results tend to be gathered—

• During germination (as in the example in this chapter).

• At the beginning and end of the rapid growth phase.

• At the end of the hardening phase (just before shipping).

• After outplanting (usually after the first 3 to 12 

months in the field, up to 5 years).

Stick to simple measurements and observations that 

are meaningful and relevant to your study. Depending on 

the subject of your experiment, your data collection may 

include one (or sometimes more) of the measurements 

described below.

Germination Rate and Percentages

A percentage of germination can be determined by comparing the total number of seeds planted versus the number 

of healthy germinants that emerge for each seed treatment. 

Discovering Ways To Improve Nursery Practices and Plant Quality 369

Figure 20.6—Stem diameter measurements are usually taken just above the medium line or on the stem (A). Height can be measured 

from the medium surface to the top of the growing point on the stem (not the top of the leaf) (B). Photos by Tara Luna.

A B
Germination rate is also important to monitor: sometimes 

the percentage of germination will ultimately be the same 

but one treatment may result in uniform and rapid germination while another treatment may be uneven or delayed. 

Daily or weekly measurements will capture differences in 

germination rate.

Plant Height and Stem Diameter

These measurements are useful to compare changes 

in plant development among the treatments and to previous crops described in the plant development records and 

propagation protocol. Stem diameter measurements are 

often taken about 0.25 in (0.5 cm) above the medium (figure 

20.6A). Height can be measured from the growing medium 

surface to the top of the growing point on the stem (not the 

top of the leaf) (figure 20.6B).

Shoot-to-Root Ratios

Shoot-to-root ratios are taken only periodically and 

usually only as small samples, because these measurements destroy the plants sampled. They are based on ovendry weight. Carefully remove any medium from the roots 

and dry the plant samples for 72 hours at 150 °F (66 °C). A 

convenient way to handle plants is to put them into paper 

lunch bags labeled by treatment and then place them into a 

drying oven. After the plants are dry, cut the sample at the 

place where the stem meets the roots (the root collar; often 

a change of color occurs here) and weigh the shoots and 

roots separately to calculate the ratio.

Plant Survival and Vigor

Plants can be subjectively rated at the beginning and end 

of each of the growth phases using a numeric rating system, 

370 Tropical Nursery Manual

Experiments Provide Opportunities To 

Learn More and Improve Nursery Practices

Figure 20.7 illustrates an innovative trial that resulted in a modest improvement in seed germination.

Figure 20.7—The small seeds of Guazuma ulmifolia, native to Puerto Rico and the U.S. Virgin Islands, are embedded in a 

hard, sweet smelling woody capsule (A). In a preliminary trial, standard treatments and a control treatment all resulted in less 

than 2 percent germination. Treatment with a mild acid solution caused each seed to produce a clear protective gel around itself, 

but the seeds still did not germinate. It was hypothesized that the seeds may benefit from passing through an animal’s digestive 

tract. Students at the University of the Virgin Islands weighed fruit, calculated seed numbers, and mixed the fruit in with feed for 

sheep in pens (B). Students then collected the sheep feces from the pens daily, removed debris with screens (C), soaked the sheep 

pellets in water and transferred the material to germination trays (D). Germination rates were higher than the control, but variable, and averaged 20 percent. Although the students did not completely solve the problem, they obtained images of the seeds’ 

development stages, and improved the original germination rate from 2 to 20 percent. Photos by Brian F. Daley.

B

A C

D
such as 0 to 5. Clear guidelines must be developed for the 

numerical scale to give a consistent relative estimate of plant 

vigor. For example, 0 = dead; 1 = no vigor, plant appears on 

verge of death; 2 = poor and slow growth; 3 = some growth, 

some vigor; 4 = plant looking vigorous; 5 = plant appears to 

be thriving and very vigorous.

Insect and Disease Analysis

The start and end of the rapid growth phase and the end 

of the hardening phase are good times to inspect for pests 

and disease. Some types of diseases, such as root rots, are 

easy to quantify and you can know what percent of your 

crop you lose to the disease. For other types of pests and 

diseases, the damage may be more subtle and you may only 

make qualitative observations. Samples of pests or diseases 

can be sent to the local agricultural extension office for 

identification, if necessary.

Outplanting Survival and Growth

Field performance after outplanting can also be evaluated by measuring height, stem diameter, survival, vigor, 

or other measures of interest (for example, amount of animal damage). See Chapter 17, Outplanting for details about 

ways to monitor performance after outplanting.

Step 7: Assessing, Recording, and 

Sharing Results

Keeping detailed records is a key part of successful experimentation. Entering observations and measurements into a 

computer or project journal is a very good practice (figure 

20.8). A simple tabular format is fine for most types of data 

and makes capturing and assessing the data easier.

While only one person is recording the data, others may 

contribute to subjective evaluations. Also, the person in 

charge of the research project may solicit the observations 

of other staff members and enter these observations in the 

journal as well.

Some experiments may focus on only one phase of growth, 

such as the germination phase. Many others will follow plants 

through all phases. Regardless, when the final phase is complete, it is important to assess the data and observations collected. Data must be organized to be interpreted. In many 

cases, data can be graphed to visually show differences among 

the controls. Any results should be shared with other staff and 

entered into the records. If the experiment focused on producing one species, the results need to be entered into the protocol 

notes for that species, even if no difference was observed. If a 

difference was observed, and one or more of the treatments 

resulted in better germination, survival, or quality than the 

control, the experiment needs to be repeated at least once or 

twice more to verify the results. In the interim, however, the 

new production technique can tentatively become the new 

protocol. If, after a few repetitions, the same results are found, 

the new technique can be adopted as the official protocol.

If no difference was observed, or if the control treatment 

performed the best, that is still very valuable information. 

Keep good records in the plant protocol notes of what was 

tried, even if it failed. Otherwise, someone might think to try 

it again, wasting time, resources, and energy. In addition, 

noting ideas for future experiments is an important part of 

concluding the experiment.

Consider sharing your experimental results at nursery meetings, by submitting short papers to professional publications such as Native Plants Journal, or by 

uploading the information into the Native Plant Network 

(http://www.nativeplantnetwork.org).

Requesting Assistance When 

Needed

The ability to carry out your own experiments as 

described in this chapter is an empowering tool for improving plant growth, quality, and survival in your nursery. 

Experimenting can also help save you and your clients 

money and time over the years. Sometimes, however, an 

idea arises for a much-needed experiment that is beyond 

the scope of what you are comfortable exploring by yourself. Perhaps the idea is too complex, costly, or time-consuming or requires more intensive statistical analyses. If 

you believe your idea would benefit you and other growers 

like you, discuss the problem you are facing and the ideas 

for possible solutions with an extension specialist or someone at your local university. Often, researchers are interested in conducting research to help native plant growers 

Discovering Ways To Improve Nursery Practices and Plant Quality 371

Figure 20.8—Keeping good records is a key part of successful 

experimentation. Photo by Tara Luna.
372 Tropical Nursery Manual

The final count of germinants from each replication 

of the control treatment is counted, and 26 out of 100 

seeds germinated (table 20.1). For the seeds that were 

mechanically scarified, 79 of the 100 seeds germinated 

(79 percent) but there were no germinants for seeds 

treated with hot water at the end of the 4 weeks (table 

20.1). A graph can be drawn to show cumulative germination on each day for each treatment (figure 20.9). 

For control seeds, cumulative germination increased 

at a constant rate throughout the 4 weeks of the trial 

and it appeared germination would continue to occur 

although the experiment had concluded. For seeds that 

were mechanically scarified, however, the graph indicates that most of the germinants emerged 3 to 8 days 

after sowing, with germination rates tapering off after 

the eighth day. This small trial showed that mechanical 

scarification yielded higher germination percentages 

and faster germination rates than the control. Although 

some variation existed from block to block, the differences among treatments within each block were similar 

indicating that the treatment results were not an isolated, 

chance event. As a result, the germination protocol for 

this species can be revised to include mechanical scarification as the seed treatment method.

The hot water treatment will also be noted in the 

protocol notes. Although a closely related species is 

known to respond well to hot water, the 20-sec treatment in 200 °F (93 °C) water on these seeds resulted 

in a 0-percent germination. It is likely that the 20-sec 

exposure was too long for this species—perhaps this 

species is smaller seeded than the related species 

described in the other protocol. A shorter time in the 

same temperature water or a lower water temperature could be tested to see if those treatments scarify 

the seeds effectively without harming them. A future 

experiment could include only 10 sec in 200 °F (93 °C) 

water or 20 sec in 170 °F (77 °C) water. If a large order 

for this species is received in the future, scarification 

will be very labor-intensive to do mechanically by 

hand. The notes about the hot water option, what did 

not work and what might work, will be a key piece of 

information for future discoveries.

In addition to the nursery’s records, the manager 

publishes the experimental findings on the Native 

Plant Network (http://www.nativeplantnetwork.org) 

to help other growers who are learning to produce this 

species.

Figure 20.9—After the final germination count, a graph can be 

drawn from the table to show cumulative germination on each 

day. Illustration by Jim Marin.

Table 20.1—Example experiment results.

Treatment Seeds treated Germinated

Control

Block 1 25 8

Block 2 25 6

Block 3 25 5

Block 4 25 7

Total 100 26 (26%)

Mechanical nick

Block 1 25 22

Block 2 25 19

Block 3 25 16

Block 4 25 21

Total 100 79 (79%)

Hot water

Block 1 25 0

Block 2 25 0

Block 3 25 0

Block 4 25 0

Total 100 0 (0%)

Example: Step 7—Assessing, Recording, and Sharing Results
and may be glad to receive suggestions from a practitioner 

about what would be a useful study. For example, the subirrigation techniques for tropical plants described in Chapter 11, Water Quality and Irrigation, came about after a 

nursery manager discussed the challenges and drawbacks 

of overhead irrigation of broad-leaved tropical plants with 

a Forest Service nursery specialist. The grower suggested 

that it would be useful to have a way to water plants from 

below instead of from above to improve water-use efficiency for tropical nurseries. The nursery specialist was 

aware of subirrigation practices, but not of applications 

for tropical seedlings, and so he passed the idea along to 

researchers at Purdue University and in the Forest Service. 

The researchers developed and tested some uses of subirrigation with tropical tree seedlings (figure 20.10). The 

methods are promising and are being shared at nursery and 

native plant conferences and in publications (Dumroese 

and others 2007). This type of cooperation between practitioners and researchers benefits growers and researchers 

and their nursery plants. Because many researchers work 

to serve the public interest, make sure you mention how the 

study would be helpful not only for your nursery, but also 

for other growers of native plants.

Field Testing To Refine the 

Target Plant

Beyond experiments to improve nursery production, 

field testing, and improving the target plant criteria for 

various sites and objectives is an important part of nursery management. This testing was described in Chapter 

3, Defining the Target Plant. At the beginning of a planting project, the land manager and the nursery manager 

agree on certain target morphological and physiological 

specifications for the plants the nursery will grow. These 

specifications are based on the eight-step assessment of 

site conditions, limiting factors, outplanting windows, 

and so on. After this prototype target plant is grown in 

the nursery, it is outplanted and monitored for survival 

and growth to assess its suitability for the site conditions. 

Both the nursery manager and the land manager learn 

from field monitoring (figure 20.11).

Discovering Ways To Improve Nursery Practices and Plant Quality 373

Figure 20.11—Field testing and refining the target plant 

criteria for various sites and objectives is an important part of 

nursery management. Illustration by Jim Marin.

Figure 20.10—If an idea for an experiment is beyond the scope 

of what you are comfortable exploring yourself, get in touch with 

an extension agent or researcher. A nursery manager discussed 

the drawbacks of overhead irrigation of broad-leaved tropical 

plants—for example, these mahogany seedlings in Palau (A)—

with a Forest Service nursery specialist. The specialist communicated the issue to researchers, who took up the challenge of testing and developing subirrigation systems for tropical nurseries 

(B). Photo A by Tara Luna and photo B by Douglass F. Jacobs.

B

A
Plots need to be monitored during the first month 

or two after outplanting and again at the end of the first 

year for initial survival (figure 20.12). Subsequent checks 

after 3 and 5 years will give a good indication of plant 

growth and survival rates. The client and the nursery 

manager then use this performance information to refine 

the target plant specifications for the next crop. The client may also alter their outplanting practices to achieve 

better survival and growth based on this information.

Beyond Experiments: Learning 

Through Reflective Practice

Managing a nursery is a profession in which you will 

learn as much or more from your own experience as you 

will learn from books, classes, and other formal teaching. 

In addition, managing a nursery is both an art and a science. Much of this book has been about the science. This 

chapter’s focus has been on learning from, and experimenting with, plants. Some types of learning, however, cannot 

come from controlled experiments; some learning comes 

from experience and reflection. The social, economic, and 

ecological context in which your nursery operates will 

change and shift over the years; your own motivations, 

values, and strategies will also evolve as you grow personally and professionally. Making room in your schedule to 

reflect on larger experiences, observations, and questions 

allows you to learn from and adapt to these inner and outer 

changes; this is called “Reflective Practice” (Schön 1983).

Reflective practice is gaining use within many professions 

where learning from direct experience is as important as formal learning; the educational and medical professions are two 

examples (Wikipedia 2011). Reflective practice is valuable for 

nursery management and other environmental professions, 

providing time and space to question and refine beliefs, values, strategies, and practices and to reflect on the changes 

happening locally and globally. Reflective time enables the 

practitioner to improve results and to align work more fully 

over time with personal values and interests.

The question of how to continually learn from experience is an important one for nursery managers. Reading 

on the subject of reflective practice or taking a training can 

help you cultivate a reflective practice. The most important 

thing is to simply be aware of the value of reflection and 

make time to reflect regularly. Setting aside even 1 hour 

per week or month for uninterrupted reflection time can 

be invaluable. Taking this time can actually save time in the 

374 Tropical Nursery Manual

Figure 20.12—The most useful measurements and observations after outplanting include plant growth rates (A, Cedrela odorata

in Costa Rica) and survival (B, Acacia koa in Hawai‘i). Photos by Douglass F. Jacobs.

A B
Discovering Ways To Improve Nursery Practices and Plant Quality 375

With the different hats I wear at the nursery and in 

my community, I am busy. But I know the time I take 

to sit and reflect has helped me to adapt to the changing 

needs of the land and people on this island. Here are 

some ways I have adapted over the years:

I noticed that when many people came to visit the 

nursery, they wished to connect with the serene environment and the forest nearby, to simply enjoy the 

land and see native plants growing. I’m probably not 

the only nursery manager in the tropics who’s had 

people come to visit and want to stay all day! I also 

enjoy sharing with my visitors. So, the question I contemplated was, how to share this environment yet still 

make a living? After a while, I found my solution. Right 

on the nursery grounds, I set up a retreat cottage for 

overnight stays, stocking it with all the creature comforts including local Kona coffee, plus field guides 

about our native birds and forests. I also began to offer 

guided forest walks to guests. This cottage has been a 

source of side income and given me a chance to share 

with visitors a deeper experience of the forest and the 

native plants growing in the nursery.

Most local restoration efforts 20 years ago, including 

mine, focused on our upland and cloud forests. Somehow we as a society were blind to the critical value of our 

endangered tropical dryland ecosystems. As the importance of dryland forests came on our radar, I did my 

research and learned ways to help restore these forests 

and to grow their plants. I now provide endemic dryland 

species including mamane, ulei, naio, lama, aweoweo, 

and many others that were not available commercially a 

decade ago. I also took on coordinating the Native Hawaiian Seed Bank Cooperative to help protect these and other 

native species by saving seed. This shift has been a good 

match between society’s changing priorities, my conservation interests, and the needs of the land.

Although my passion is growing plants, I’ve long been 

concerned about helping the native wildlife that depend 

on these plants. I have taken time over the years to sit in 

observation in the local forests and watch the ways plants 

and birds associate with each other. I know birds that eat 

the manono with dark purple fruit, pilo with orange fruit, 

akala, the Hawaiian raspberry (figure 20.13A), and naio 

with white fruit. Observing the birds taught me to prioritize learning to grow their important habitat plants in my 

nursery. The observation and reflection also helps me to 

better understand processes and interactions in a diverse 

and healthy forest, to better mimic them in restoration of 

disturbed lands.

I don’t know what the future will bring. But I know 

as I take time to reflect, learn, and stay aware of what is 

going on around me as well as within, my nursery will be 

able to evolve with the changing times.

—Jill Wagner, Forestry Consultant, Kailua Kona, HI

Owner, Future Forests Nursery, LLC

Coordinator, Hawai‘i Island Native Seed Bank Cooperative

Figure 20.13—Ripe fruits of ‘akala, the native Hawaiian 

raspberry (Rubus hawaiensis), observed to be an important 

food source for native birds (A). Jill Wagner at a dryland 

forest outplanting site (B). Photo A by J.B. Friday, and photo 

B by Yvonne Yarber Carter.

B

A

Learning and Adapting Through Reflective Practice
long run by raising awareness of inefficiencies and helping 

prioritize where to focus energy and resources. Examples of 

reflective practices include—

• Keeping a journal and reviewing it from time to time.

• Revisiting your nursery’s vision statement, guiding 

principles, roles, and goals.

• Interacting with clients a few years after planting to 

follow up on the experience.

• Contemplating your own practices and beliefs, the 

results you are currently achieving, and your own 

dreams or ideas.

• Considering the changing community needs and public concerns, and how these criteria may align with 

your own values and the nursery’s vision.

• Sitting and observing interactions in healthy ecosystems similar to the ecosystems you work to restore, 

and making notes of what you learned.

• Reviewing photos of your plants, projects, and nursery 

operations and reflecting on your thoughts and feelings.

• Taking time after a meeting or conference to sum up 

what you learned and what it might mean for you and 

your work.

• Sharing your insights with colleagues or friends.

Whether your nursery is large or small, for-profit, nonprofit, or public, reflective practice will help you continually 

learn, adapt, and evolve.

References

Dumroese, R.K.; Jacobs, D.F.; Davis, A.S.; Landis, T.D. 2007. An 

introduction to subirrigation in forest and conservation nurseries 

and some preliminary results of demonstrations. In: Riley, L.E.; 

Dumroese, R.K.; Landis, T.D., tech. coords. 2007. National proceedings: forest and conservation nursery associations—2006. 

Proc. RMRS-P-50. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 20-26.

Dumroese, R.K.; Wenny, D.L. 2003. Installing a practical research 

project and interpreting research results. Tree Planters’ Notes. 

50(1): 18–22.

Dumroese, R.K.; Luna, T.; Landis, T.D. 2008. Nursery manual for 

native plants: volume 1, a guide for tribal nurseries. Agriculture 

Handbook 730. Washington, DC: U.S. Department of Agriculture, Forest Service. 302 p. 

Lilleeng-Rosenberger, K.E. 2005. Growing Hawai‘i’s native plants: 

a simple step-by-step approach for every species. Honolulu: Mutual 

Publishing. 420 p.

Sagan, C. 1996. The demon-haunted world: science as a candle in 

the dark. New York: Ballantine Books: 480 p.

Schön D. 1983. The reflective practitioner: how professionals 

think in action. New York: Basic Books.

Wikipedia. 2011. Reflective practice. http://en.wikipedia.org/

wiki/Reflective_practice. (November 2011).

Additional Reading

Landis, T.D.; Tinus, R.W.; McDonald, S.E.; Barnett, J.P. 1994. 

The container tree nursery manual: volume 1, nursery planning, 

development, and management. Agriculture Handbook 674. 

Washington, DC: U.S. Department of Agriculture, Forest Service. 

188 p.

White, T. L. 1984. Designing nursery experiments. In Duryea, 

M.L.; Landis T.D. (eds.). 1984. Forest nursery manual: production of bareroot seedlings. Martinus Nijhoff/Dr W. Junk 

Publishers. The Hague/Boston/Lancaster, for Forest Research 

Laboratory. Corvallis: Oregon State University: 291-306.

376 Tropical Nursery Manual

Tropical Nursery Manual

Tropical ecosystems and agroecosystems are vital, life-giving landscapes. They are 

home to diverse plants, animals, and people in a range of climatic, geologic, cultural, and 

environmental contexts. Protecting and revitalizing these systems involves planting native 

and traditional species, and sowing awareness of these plants among people. Tropical 

nurseries are a key link in restoring tropical ecosystems and agroecosystems, and can 

integrate goals of economic resiliency, cultural renewal, and more.

This Tropical Nursery Manual serves people who are starting or operating a nursery 

for native and traditional species in the tropics. Key concepts, principles, and processes 

are presented, based on proven practices and the best science available. Understanding 

these concepts and principles will make it easier to operate a nursery successfully, and 

to meet project objectives in the field. Topics covered include nursery planning, plant 

propagation, crop production, plant care, outplanting, and ongoing learning.

This manual also recognizes that every nursery is unique. Local conditions and 

ingenuity, integrated with the information in this manual, combine to cultivate highquality plants with the best chance to survive and flourish into the future.

“A very useful and urgently needed tool for native plant growers and nursery 

managers in the tropics. It highlights the challenges faced when dealing with tropical 

species no one else has propagated.”

— Christian Torres-Santana

USDA Forest Service, International Institute of Tropical Forestry, 

San Juan, Puerto Rico

“This manual not only has great, detailed information on how to manage the 

small-scale, highly diverse nurseries for native and traditional tropical plants, but 

it also includes inspirational case studies from nursery practitioners working in 

the field today.”

— J.B. Friday

Tropical Forestry & Agroforestry Extension, University of Hawai‘i, Hilo, Hawai‘i

 “The Tropical Nursery Manual is very complete, covering thoroughly all aspects of 

nursery techniques in the tropics. But, it also provides clear ideas about how to solve 

arising problems. The attractive presentation with the excellent visual material 

guarantees an easy understanding for practical application. I am sure that this 

manual sets a new standard in the subject of nursery management in the tropics 

and will accompany many nursery managers and foresters in the years to come.”

— Laszlo Pancel

GIZ Advisor, Editor, Tropical Forestry Handbook Editions I & II, 

La Libertad, El Salvador

“This manual is a wonderful resource for people who want to do the best job they 

can growing tropical trees. The authors provide helpful information, taking into 

account many types of situations and giving the reader ideas about how to solve 

specific issues, as well as the idea that one can be innovative and find solutions to 

meet any challenge.”

— Jill Wagner

Future Forests Nursery, Kailua-Kona, Hawai‘i