Diseases of Trees in the Great Plains

United States Department of Agriculture

Forest Rocky Mountain General Technical

Service Research Station Report RMRS-GTR-335 November 2016
To learn more about RMRS publications or search our online titles:

www.fs.fed.us/rm/publications

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Bergdahl, Aaron D.; Hill, Alison, tech. coords. 2016. Diseases of trees in the Great Plains. 

Gen. Tech. Rep. RMRS-GTR-335. Fort Collins, CO: U.S. Department of Agriculture, 

Forest Service, Rocky Mountain Research Station. 229 p. 

Abstract 

Hosts, distribution, symptoms and signs, disease cycle, and management strategies are 

described for 84 hardwood and 32 conifer diseases in 56 chapters. Color illustrations are 

provided to aid in accurate diagnosis. A glossary of technical terms and indexes to hosts 

and pathogens also are included. 

Keywords: Tree diseases, forest pathology, Great Plains, forest and tree health, 

windbreaks.

Cover photos by: James A. Walla (top left), Laurie J. Stepanek (top 

right), David Leatherman (middle left), Aaron D. Bergdahl (middle right), 

James T. Blodgett (bottom left) and Laurie J. Stepanek (bottom right).
Background

This technical report provides a guide to assist arborists, landowners, woody plant 

pest management specialists, foresters, and plant pathologists in the diagnosis and 

control of tree diseases encountered in the Great Plains. It contains 56 chapters on tree 

diseases prepared by 27 authors, and emphasizes disease situations as observed in the 

10 states of the Great Plains: Colorado, Kansas, Montana, Nebraska, New Mexico, 

North Dakota, Oklahoma, South Dakota, Texas, and Wyoming. 

The need for an updated tree disease guide for the Great Plains has been recognized for some time and an account of the history of this publication is provided here. 

When the Pest Management Task Force of the Great Plains Agricultural Council, 

Forestry Committee, met in 1980, the task force requested that the committee support 

its publication of a tree disease handbook for the Great Plains, and assist it in obtaining the necessary funding. The request was approved. 

During the task force’s 1981 meeting, a working group was formed to prepare a 

prospectus for the handbook. The working group members, Edward Sharon, James 

Walla, Mark Harrell, and Jerry Riffle, chose specific diseases for inclusion in the 

handbook, selected potential authors for each chapter and determined chapter format 

to emphasize diagnosis, biology, damage, and management of diseases. In 1982, 

the Pest Management Task Force agreed that Riffle would serve as coordinator of 

the handbook, with Glenn Peterson as co-coordinator. Riffle contacted all potential 

contributors in May 1982, and their response was excellent; 31 persons agreed to be 

authors or coauthors of chapters on 64 diseases. The source of funding for publication of the handbook was resolved in 1985 when it was proposed that it be published 

as a general technical report by the U.S. Department of Agriculture (USDA) Forest 

Service, Rocky Mountain Research Station (RMRS).

This proposal was approved by the Executive Committee of the Great Plains 

Agricultural Council, Forestry Committee. The original Diseases of Trees in the Great 

Plains1, published in 1986, has been an extremely popular publication used by a wide 

spectrum of natural resource professionals in the Great Plains and beyond.

During the Great Plains Tree Pest Council annual meeting in 2012, a discussion 

was initiated about updating the original publication to include new diseases and to 

reflect the current available knowledge about tree diseases in the Great Plains. Upon 

agreement to the update, a group was formed consisting of James Walla, William 

Jacobi, Mark Harrell, Judith O’Mara, John Ball, James Blodgett, Jared LeBoldus, 

Simeon Wright, Laurie Stepanek, Les Koch, Rachel Allison, Nicole Ricci, and 

Aaron Bergdahl to consider which new diseases to include and potential authors and 

reviewers. Letters of support for the effort were requested from the state foresters of 

the Great Plains states and subsequently approval of the update was given by RMRS 

Station Director Sam Foster. Possible funding sources were researched, and Michele 

Schoeneberger, from the USDA National Agroforestry Center, offered assistance and 

set aside funds for printing the publication. Later, Roy Mask of USFS Region 2 came 

forward with additional funding in 2015 and Gregg DeNitto and Marcus Jackson of 

Northern Region Forest Health Protection facilitated funding in 2016.

The chair of the Great Plains Tree Pest Council at that time, Aaron Bergdahl, and 

Alison Hill of the RMRS offered to organize the update effort and serve as technical 

editors. Work on the update began by requesting the contribution of 27 authors with 

forest pathology experience in the Great Plains. The response from the authors was 

very positive. The chapter content and technical reviews were completed in late 2013 

and 2014. The updated publication, which is available in print and online versions, is 

the culmination of the cooperative efforts of numerous professionals dedicated to the 

health of forest resources in the Great Plains from several state and federal agencies 

and research institutions.

1 Riffle, Jerry W.; Peterson, Glenn W., tech. coords. 1986. Diseases of trees in the Great 

Plains. Gen. Tech. Rep. RM-129. Fort Collins, CO: U.S. Department of Agriculture, Forest 

Service, Rocky Mountain Forest and Range Experiment Station. 149 p.

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Dedication

This update of Diseases of Trees in the Great Plains is dedicated to the coordinators of the original manuscript, Jerry Riffle and Glenn Peterson. These two retired 

U.S. Forest Service researchers are to be commended for their many years of dedication 

and contributions to the understanding of tree diseases and health in the Great Plains 

region. Their passion for their field of study and Great Plains forest resources was embodied in the first edition of this valuable guide. The authors of this update have done 

their utmost to carry on the traditions of scientific inquiry and excellence personified 

by Riffle and Peterson in Great Plains forest pathology.

Authors and Their Affiliations

Gerard C. Adams Associate Professor of Practice, Department of Plant 

Pathology, University of Nebraska, Lincoln, NE 68583

Rachel A. Allison Western Forest Health Specialist, Nebraska Forest Service, 

University of Nebraska, North Platte, NE 69101

Jon A. Appel Plant Pathologist, Kansas Department of Agriculture, 

Manhattan, KS 66503

David N. Appel Plant Pathologist, Texas A&M University, College Station, 

Texas 77843

John Ball Forest Health Specialist, South Dakota Department of 

Agriculture and Forestry Extension, South Dakota State 

University, Brookings, SD 57007

Aaron D. Bergdahl Forest Health Manager, North Dakota Forest Service, North 

Dakota State University, Fargo, ND 58102

James T. Blodgett Plant Pathologist, U.S. Forest Service, Forest Health 

Protection, Rapid City, SD 57702

Kelly S. Burns Plant Pathologist, U.S. Forest Service, Rocky Mountain 

Region, State and Private Forestry, Golden, CO 80401

Whitney Cranshaw Entomologist, Colorado State University, Fort Collins, CO 

80523

Kelsey L. Dunnell Research Specialist, Department of Plant Pathology, North 

Dakota State University, Fargo, ND 58102

Mark L. Gleason Professor, Department of Plant Pathology and Microbiology, 

Iowa State University, Ames, IA 50011

Mark O. Harrell Forest Health Program Leader, Nebraska Forest Service, 

University of Nebraska, Lincoln, NE 68583

Thomas C. Harrington Forest Pathologist, Department of Plant Pathology and 

Microbiology, Iowa State University, Ames, IA 50011

Alison Hill Research Program Manager, Forest and Woodlands 

Ecosystems Science Program, Rocky Mountain Research 

Station, Flagstaff, AZ 86001 

Marcus B. Jackson Plant Pathologist, U.S. Forest Service, Forest Health 

Protection, Missoula, MT 59807

William R. Jacobi Professor of Forest and Shade Tree Pathology, Department 

of Bioagricultural Sciences and Pest Management, 

Colorado State University, Fort Collins, CO 80523 

(retired)

Jennifer Juzwik Research Plant Pathologist, U.S. Forest Service, Northern 

Research Station, St. Paul, MN 55108
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Megan M. Kennelly Associate Professor and Extension Specialist, Department of 

Plant Pathology, Kansas State University, Manhattan, KS 

66506

Les Koch Forest Health Specialist, Wyoming State Forestry Division, 

Cheyenne, WY 82002

Jared M. LeBoldus Assistant Professor, Department of Botany and Plant Pathology/ 

Forest Engineering and Resources Management, Oregon 

State University, Corvallis OR, 97331

Phillip A. Mason National Operations Manager, USDA Animal and Plant 

Health Inspection Service, Plant Protection and 

Quarantine, Fort Collins, CO 80526

Douglas L. McNew Mycologist, Department of Plant Pathology and 

Microbiology, Iowa State University, Ames, IA 50011

Judith O’Mara Diagnostician, Department of Plant Pathology, Kansas State 

University, Manhattan, KS 66506

Hayley M. Nelson Research Associate II, Dept. of Plant Pathology and 

Microbiology, Iowa State University, Ames, IA 50011

Nicole Opbroek Forest Health Specialist, Kansas State Forest Service, 

Manhattan, KS 66506

Michael E. Ostry Research Plant Pathologist, U.S. Forest Service, Northern 

Research Station, St. Paul, MN 55108

Scott C. Redlin Plant Pathologist, USDA APHIS Plant Protection and 

Quarantine Center for Plant Health Science and 

Technology Raleigh, NC 27606-5202 

Glen R. Stanosz Professor, Department of Forest and Wildlife Ecology, 

University of Wisconsin–Madison, Madison, WI 53706

Laurie J. Stepanek Forest Health Specialist, Nebraska Forest Service, 

University of Nebraska, Lincoln, NE 68583

Ned Tisserat Extension Specialist and Professor, Colorado State 

University, Fort Collins, CO 80523 (retired)

Timothy C. Todd Research Nematologist, Kansas State University, Manhattan, 

Kansas 66506

James A. Walla Northern Tree Specialties Fargo, ND 58102

James J. Worrall Forest Pathologist, U.S. Forest Service, Rocky Mountain 

Region, Forest Health Protection, Gunnison, CO 81230

Simeon Wright Forest Pathologist, Missouri Department of Conservation, 

Columbia, MO 65201

Acknowledgments

Special thanks to Sam Foster, former director of the Rocky Mountain Research Station; 

the National Agroforestry Center’s Michele Schoeneberger; and the state foresters of the 

Great Plains states for their support of the update of this guide. We also thank the many 

professionals who critically reviewed individual manuscripts for accuracy and suggested 

improvements. All contributions to the content and images in this guide are gratefully 

acknowledged. In particular, James A. Walla is recognized for his dedicated technical 

work reviewing this guide, providing valuable advice and crucial support of this update. 

Laurie J. Stepanek is also acknowledged for her extensive technical review and support 

of this effort. Gerard C. Adams is thanked for providing a thorough review of fungal 

nomenclature. Finally, thanks go to all the authors who cooperated across agencies and 

institutions and completed the excellent update of this valued tree disease guide.
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Contents

Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . i

Dedication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ii

Authors and Their Affiliations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ii

Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . iii

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1. Environmental Stresses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

John Ball; revised from Donald F. Schoeneweiss 

(Riffle and Peterson 1986)

2. Anthracnose Diseases of Broadleaf Trees . . . . . . . . . . . . . . . . . . . . . . . . . 12

Scott C. Redlin; revised from Robert W. Stack and 

Kenneth E. Conway (Riffle and Peterson 1986)

3. Bur Oak Blight . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

Thomas C. Harrington and Douglas L. McNew

4. Cherry Leaf Spots . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20

Gerry Adams, Ned Tisserat, and William Jacobi

5. Chlorosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26

Laurie J. Stepanek, Mark O. Harrell, and Rachel A. Allison; 

revised from Mark O. Harrell and Mark W. Andrews (Riffle and 

Peterson 1986)

6. Herbicide Damage to Trees and Shrubs . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

Aaron Bergdahl and William Jacobi

7. Taphrina Diseases of Shade and Fruit Trees . . . . . . . . . . . . . . . . . . . . . . . 35

Mark L. Gleason, Hayley M. Nelson, and Megan M. Kennelly

8. Leaf Tatter and Leaf Tatters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

Laurie J. Stepanek and Mark O. Harrell

9. Marssonina Leaf Spot and Blight of Populus Species . . . . . . . . . . . . . . . . . 41

Michael E. Ostry and Jared M. LeBoldus; revised from 

John E. Watkins and David S. Wysong (Riffle and Peterson 1986)

10. Melampsora Leaf Rust of Cottonwood, Aspen, and Willow . . . . . . . . . . . . 44

Phillip A. Mason and Jared LeBoldus; revised from Glenn W. 

Peterson and Robert W. Stack (Riffle and Peterson, 1986)

11. Mycosphaerella Leaf Spot of Ash . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46

Judith O’Mara and Megan Kennelly

12. Powdery Mildew of Lilac . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48

John Ball; revised from Richard Dorset and 

Michael W. Ferguson (Riffle and Peterson, 1986)

13. Tar Spots of Maple . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51

Aaron D. Bergdahl and Kelsey L. Dunnell

14. X-Disease of Chokecherry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55

James A. Walla; revised from Glenn W. Peterson and 

David W. Johnson (Riffle and Peterson 1986)

15. Biscogniauxia Canker of Oak . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58

Judy O’Mara and Nicole Opbroek

16. Botryodiplodia Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61

Mark O. Harrell and Laurie J. Stepanek; revised from 

Jerry W. Riffle and Joseph M. Krupinsky, Botryodiplodia canker 

of elms; Glenn W. Peterson and Harrison L. Morton, Botryodiplodia 

disease of Russian-olive; and Robert Lewis, Jr., and Kenneth 

E. Conway, Botryodiplodia canker of sycamore (Riffle and 

Peterson 1986)

17. Brown Rot of Stone Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

John Ball and James A. Walla; revised from David S. Wysong 

and James A. Walla (Riffle and Peterson 1986)

18. Black Knot of Cherry and Plum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67

Mark O. Harrell and James T. Blodgett; revised from 

David S. Wysong and Mark O. Harrell (Riffle and Peterson 1986)
19. Black-Spot Nectria Canker . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70

Marcus B. Jackson; revised from Tubercularia Canker of 

Siberian Elm and Russian-olive, by Joseph M. Krupinsky and 

James A. Walla (Riffle and Peterson 1986)

20. Canker and Dieback of Russian-Olive . . . . . . . . . . . . . . . . . . . . . . . . . . . 74

Gerry Adams, Ned Tisserat, and William Jacobi

21. Cankers and Root Collar Rot of Honeylocusts . . . . . . . . . . . . . . . . . . . . . 78

William Jacobi and James A. Walla

22. Crown Gall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84

Simeon Wright; revised from Michael W. Ferguson and 

Richard Dorset

23. Cryptodiaporthe (Dothichiza) Canker of Populus Species . . . . . . . . . . . 86

Michael E. Ostry and Jared M. LeBoldus; revised from David W. 

Johnson and Robert W. Stack (Riffle and Peterson 1986)

24. Cryptosphaeria Canker of Cottonwood and Aspen . . . . . . . . . . . . . . . . . . 88

James J. Worrall, Kelly S. Burns, and Jennifer Juzwik; 

revised from Thomas E. Hinds and Jerry W. Riffle (Riffle and 

Peterson 1986)

25. Cytospora Canker of Hardwoods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

Gerard Adams and William Jacobi

26. Fire Blight of Apple, Pear, and Other Ornamental Rosaceous 

Shrubs and Trees . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94

Megan M. Kennelly and Mark L. Gleason

27. Hypoxylon Canker of Aspen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97

Michael E. Ostry and Jared M. LeBoldus; revised from 

Thomas E. Hinds and Jerry W. Riffle (Riffle and Peterson 1986)

28. Phomopsis Canker of Poplar . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100

Michael E. Ostry and Jared M. LeBoldus; revised from: 

Theodore H. Filer and Edward M. Sharon (Riffle and Peterson 1986)

29. Septoria Leaf Spot and Canker of Poplar . . . . . . . . . . . . . . . . . . . . . . . . 102

Jared M. LeBoldus, Michael E. Ostry, and James T. Blodgett; 

revised from Joseph M. Krupinsky, David W. Johnson, 

Jerry W. Riffle, and David S. Wysong (Riffle and Peterson 1986)

30. Stem Decays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105

James J. Worrall

31. Thousand Cankers Disease (TCD) of Walnut . . . . . . . . . . . . . . . . . . . . . 118

Ned Tisserat and Whitney Cranshaw

32. Wetwood (Slime Flux) and Alcohol Flux in Hardwoods . . . . . . . . . . . . . . 120

William Jacobi and John Ball

33. Witches’-Broom of Hackberry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124

Mark O. Harrell and Laurie J. Stepanek; revised from 

Mark O. Harrell and Frederick J. Crowe (Riffle and 

Peterson 1986)

34. Dutch Elm Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126

Kelsey L. Dunnell and Aaron Bergdahl

35. Oak Wilt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129

Jennifer Juzwik and David N. Appel

36. Verticillium Wilt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134

Mark O. Harrell and Laurie J. Stepanek; revised from 

David S. Wysong and Mark O. Harrell (Riffle and 

Peterson 1986)

37. Yellows Disease of Ash, Lilac, and Elm . . . . . . . . . . . . . . . . . . . . . . . . . . 137

James A. Walla and Leslie Koch; partially revised from 

Wayne A. Sinclair and David S.Wysong (Riffle and 

Peterson 1986)

38. Armillaria Root Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 141

James T. Blodgett; revised from Lloyd R. Fuller and 

Robert L. James (Riffle and Peterson 1986)

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39. Ganoderma Root Rot or White Mottled Rot . . . . . . . . . . . . . . . . . . . . . . 145

James T. Blodgett

40. Nematodes of Broadleaf Trees . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 148

Timothy C. Todd and Jon Appel

41. Brown Spot of Pine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152

Judith O’Mara and Aaron Bergdahl

42. Cercospora Blight of Junipers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155

Glenn Peterson and David Wysong (Riffle and Peterson 1986) 

(updated by Ned Tisserat)

43. Dothistroma Needle Blight of Pines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157

Aaron Bergdahl and Judy O’Mara; revised from Glenn W. 

Peterson and David Wysong (Riffle and Peterson 1986)

44. Kabatina Blight of Juniper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161

Andrea Ostrofsky and Glenn Peterson (Riffle and Peterson 

1986) (updated by Ned Tisserat)

45. Lirula Needle Blight of Spruce . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163

James A. Walla

46. Cyclaneusma Needle Cast of Pines . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166

James A. Walla; revised from Glenn W. Peterson and 

James A. Walla (Riffle and Peterson 1986)

47. Phomopsis Blight of Junipers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169

Glenn Peterson (Riffle and Peterson 1986) 

(updated by Ned Tisserat)

48. Rhizosphaera Needle Cast of Spruce . . . . . . . . . . . . . . . . . . . . . . . . . . . 171

James A. Walla; revised from Darroll D. Skilling and 

James A. Walla (Riffle and Peterson 1986)

49. Stigmina Needle Cast of Spruce . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174

James A. Walla and Aaron D. Bergdahl

50. Canker Diseases of Juniper and Oriental Arborvitae . . . . . . . . . . . . . . . 177

Ned Tisserat

51. Diplodia Shoot Blight and Canker Disease . . . . . . . . . . . . . . . . . . . . . . . 180

James T. Blodgett and Glen R. Stanosz; revised from 

Glenn W. Peterson and David W. Johnson (Riffle and 

Peterson 1986)

52. Gymnosporangium Rusts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184

Simeon Wright, James A. Walla, and William R. Jacobi; 

revised from James A. Walla and Jerry Riffle and from 

Kenneth E. Conway and Mark W. Andrews (Riffle and 

Peterson 1986)

53. Valsa (Cytospora) Canker of Spruce . . . . . . . . . . . . . . . . . . . . . . . . . . . . 190

James A. Walla and Aaron D. Bergdahl; revised from 

James A. Walla and Frederick J. Crowe (Riffle and Peterson 1986)

54. Western Gall Rust of Pines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193

James T. Blodgett and Kelly S. Burns; revised from 

Glenn W. Peterson and James A. Walla (Riffle and Peterson 1986)

55. Pine Wilt Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196

Timothy C. Todd and Mark Harrell

56. Root Parasitic Nematodes in Junipers and Pines . . . . . . . . . . . . . . . . . . 200

Timothy C. Todd and Jon Appel; revised from Glenn W. Peterson 

and Jerry W. Riffle (Riffle and Peterson 1986)

Glossary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 203

Index To Host Plants With Scientific Equivalents . . . . . . . . . . . . . . . . . . . . . 211

Index to Plant Diseases and Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 

Index to Associated Insects, Nematodes, and Pathogen Vectors . . . . . . . . 227

Index to Associated Chemical Article Number . . . . . . . . . . . . . . . . . . . . . . . . 228
viii
USDA Forest Service RMRS-GTR-335. 2016. 1

Introduction

Scope

The purpose of this handbook is to assist users in diagnosing tree diseases and reducing their impact. Included are the most damaging diseases, along with some that are the 

subject of frequent inquiry although of minor importance. The diseases included were 

those encountered primarily on trees in conservation plantings such as windbreaks, 

farmstead plantings, and living snow fences and in landscape plantings in recreational 

areas and near rights-of-way. Additionally, diseases affecting tree nurseries and Christmas 

tree farms in the Great Plains are included. By association, there is great overlap with the 

most common diseases encountered in native woodlands and urban plantings.

Diseases of fruit and nut trees are included when they are encountered on trees that are 

used in the above types of plantings. Information on nutrient deficiencies is not included, 

except in the chapter on chlorosis. There is no extensive coverage of damage by airborne 

pollutants except that caused by herbicides. The chapters are grouped as hardwood 

diseases and conifer diseases; within each of these categories they are classified as to the 

part of plant affected: foliage, branch and stem, vascular wilt, and root and soilborne. 

Most of the diseases are treated separately in the following standard format: “Hosts and 

Distribution,” “Symptoms and Signs,” “Disease Cycle,” “Damage,” “Management,” 

and “Selected References.” Select chapters cover several similar diseases affecting a 

particular host or similar types of pathogens, as in the chapter on stem decays (chapter 

30). The emphasis of this tree disease guide is on disease situations in the Great Plains. 

Accordingly, no special effort has been made to include the total distribution of pathogens, or to list all of their hosts. Similarly, “Symptoms and Signs” and “Disease Cycles” 

are described primarily as observed in the Great Plains.

Diagnosis

In making a diagnosis, the reader is referred first to the host index to see which diseases are listed for the tree host in question. Then the reader should check the writeup on 

the diseases for that host and also the photographs to see if they conform to the disease 

under consideration. A glossary of technical terms is included for a more complete understanding of the chapters. English units of measure are used for the most part; metric units 

are used primarily in descriptions of pathogens.

Management

Information on management (reducing damage) is provided. Where chemicals are suggested or recommended, the reader must determine if the material is currently registered 

for the intended use in the area of application. Most of the articles listed in “Selected 

References” were chosen because they will be useful in understanding the recommendations for reducing damage. Because information on the cycle of some diseases is limited, 

the recommendations for their control are in some cases quite general.

Damage from pathogens can often be reduced by increasing the vigor of trees. Thus, 

the “Management” section of several of the chapters in this guide includes recommendations for increasing vigor. Vigor of trees in the Great Plains usually can be increased by 

supplemental watering. Application of fertilizers sometimes can result in increased tree 

vigor; however, the wholesale application of fertilizers for increasing tree vigor is not 

recommended. Use of fertilizers should be based on evidence that the tree in question 

(considering species and age) will respond to fertilizers on the site involved.

History

The demand for trees in the Great Plains stems from early settlers’ needs for orchards, 

woodlots, windbreaks, and hedges. Long before the Federal Timber Culture Act of 1873, 

individuals, companies, associations, and local governments had promoted tree planting in the Great Plains. Establishment of dry-land Agricultural Experiment Stations at 

Mandan, North Dakota; Woodward, Oklahoma; and Cheyenne, Wyoming; resulted in 

increased planting of trees by farmers and ranchers, who cooperated with these stations 

in tests of tree species, cultural methods, spacing of trees, and the number of rows needed 
2 USDA Forest Service RMRS-GTR-335. 2016.

(Droze 1977). Their work aided the USDA’s tree distribution program authorized by 

the Clarke-McNary Act, which began in 1924. This program, under the direction of the 

Extension Service of the USDA, encouraged tree planting in the Great Plains. 

The biggest impetus to tree planting came, however, when President Franklin D. 

Roosevelt, by executive order in July 1934, authorized the Shelterbelt Project. This 

project, later called the Prairie States Forestry Project, operated between 1935 and 1942. 

It led to the planting of 223 million trees in 18,600 miles of field windbreaks. Tree planting for protection purposes continued after 1942 under the direction of the USDA Soil 

Conservation Service. Tree planting was not as extensive as during the Prairie States 

Forestry Project, but was fairly steady. The Soil Bank program of the 1960s resulted in an 

increase in plains tree planting as considerable land was taken out of crop production and 

planted to trees.

Rising energy costs spurred tree planting for many additional years, not only on farms and 

ranches but in communities as well. Trees were also planted along roads for “living snow 

fences.” In more recent years, tree planting in communities has continued, but rural tree plant-

ing has declined. Furthermore, strong crop prices and larger farm machinery have led to the 

loss of shelterbelts, with farmers opting to remove older shelterbelts rather than renovating or 

replanting.

Although many pest problems were noted in the early plantings in the Great Plains, 

very little systematic study of tree diseases and insects was conducted, with the exception of diseases of seedlings. Research on diseases of tree seedlings in the Great Plains 

was initiated some 100 years ago by Carl Hartley and USDA colleagues. Their research 

resulted in a good understanding of damping-off disease of pines and other conifers and 

provided some information on other diseases of conifer seedlings.

Little additional tree disease research occurred in the Great Plains until the period of 

the Prairie States Forestry Project (1935–1942). With the expansion of existing nurseries 

and the establishment of many new nurseries to provide trees for that project, a number 

of disease problems were encountered, particularly on hardwood seedlings. Thus, during 

this period, Ernest Wright and USDA colleagues concentrated their research on disease 

problems of hardwood seedlings, with some work being done on plantation diseases such 

as Phymatotrichum root rot and Cytospora canker of cottonwood (Populus deltoides).

When the Prairie States Forestry Project was ended in 1942, tree disease research in 

the Great Plains by the USDA was discontinued. Research by land grant institutions 

at this time was minimal as well. Agricultural extension personnel in the Great Plains 

accumulated some information on tree diseases, but they had little time to devote to 

tree problems. Likewise, pest management specialists assigned to U.S. Forest Service 

regional offices devoted little time to tree disease problems in the Great Plains, other than 

those associated with federal nurseries and national monuments.

 A turnaround came in 1958 when a plant pathologist position was established in the 

Lincoln Unit of the U.S. Forest Service’s Rocky Mountain Forest and Range Experiment 

Station. Nursery diseases received attention first; diseases in pine and juniper plantings 

were emphasized later. With the hiring of another pathologist in 1972, research was 

expanded to include additional research on hardwood diseases and on mycorrhizae. Land 

grant institutions in North Dakota, South Dakota, Nebraska, Kansas, and Oklahoma 

expanded their research efforts and more extension plant pathologists were hired. 

U.S. Forest Service pest management specialists increased their activities in the Great 

Plains, and pest management specialists were assigned to several state forester offices. 

Additionally, The ARS added a tree disease research project to support a tree improvement project in the Northern Plains.

The increased effort on tree diseases prompted the Great Plains Agricultural Council, 

Forestry Committee to establish a Pest Management Task Force to increase communication among workers and to coordinate activities in research, pest management, and 

extension. This task force sponsored the original development of this handbook. 

Since the publication of this guide in 1986, land grant institutions in the Great Plains 

have continued to support their capacity to study tree disease problems of concern, 

and federal programs have maintained their support of state forest health programs and 

research groups within the U.S. Forest Service. But because funding for tree disease 
USDA Forest Service RMRS-GTR-335. 2016. 3

research and forest health programs has not increased at a rate sufficient to sustain the 

number of positions and programs that existed when the original version of this guide 

was published, the future state of forest pathology knowledge in the Great Plains is at 

the moment uncertain. Further, the number of personnel in the Great Plains focusing on 

diseases of trees is decreasing and fewer positions are being replaced after retirement. 

While this guide was being updated, three senior active members of the Great Plains Tree 

Pest Council and the greater forest pathology scientific community retired. With several 

more senior researchers scheduled to retire in the coming years, the future state of forest 

pathology knowledge in the Great Plains will diminish without a committed effort to 

maintain and rebuild well-funded programs and increase pathology expertise in the region. With respect to these current challenges, updating this disease guide at this time was 

critical to fully capitalize on the wealth of expertise held by senior members of the Great 

Plains Tree Pest Council.

Use and Safe Handling of Pesticides

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 U.S. Environmental Protection Agency, consult your state department of agriculture 

to be sure the intended use is still registered. The rules below should be followed when 

handling pesticides. These rules should be read by all persons involved in pesticide use. 

Copies of these rules should be posted in several places, particularly in the areas of pesticide storage.

Pesticides are poisonous and always should be used with caution. The dangers associated with mishandling and misapplication of pesticides include possible injury to the 

operator and handler and damage to target and non-target plants, to the equipment, and to 

the environment. Read the health and safety codes of your organization pertaining to use 

of toxic chemicals prior to use of pesticides.

1. Read the label. Handlers should read, understand, and follow all instructions on 

the label. Notice warnings and cautions before opening the container. Repeat the 

procedure every time, no matter how familiar you think you are with the directions. 

Apply the material only in the amounts and at the times specified.

2. Avoid contact. Avoid inhaling sprays and dusts. Avoid contact with skin and eyes. 

When directed by the label, wear the proper protective clothing, a mask, and all 

other recommended personal protective equipment. Do not eat, drink, smoke or 

chew on anything while spraying or dusting. Wash thoroughly before eating.

3. Apply safely. Use only the specified dosages and mix as directed. Do not use your 

mouth to siphon liquids from containers or blow out clogged lines or nozzles. Use 

clean, well-functioning equipment to apply the pesticides. Do not spray with leaking 

hoses or connections. Do not work or allow others to work in the drift of the spray 

or dust.

4. Wash immediately. Stop and wash off any pesticide spilled on the body. Remove 

contaminated clothing. Wash and change to clean clothing after spraying and 

dusting. Also, wash clothing each day before re-use.

5. Dispose of containers properly. Always dispose of empty containers so that they pose 

no hazard to humans, animals, or plants (either terrestrial or aquatic). When unsure 

about proper disposal procedures, contact the nearest waste management authority.

6. Store safely. Keep pesticides stored together outside the home or office away from 

food and usual working areas. Keep them under lock and key. Label and sign the 

area well and do not store other chemicals among the pesticides. Always keep the 

pesticides in the original containers, and keep them tightly closed.

7. Report illness. If symptoms of illness occur during or shortly after dusting or spraying, 

call a physician or poison control number or 911 and get the patient to a hospital 

immediately.
4 USDA Forest Service RMRS-GTR-335. 2016.

1. Environmental Stresses

John Ball; revised from Donald F. Schoeneweiss 

(Riffle and Peterson 1986)

Environmental stresses, which are nonliving (abiotic) disorders, are commonly 

overlooked causal agents when tree problems are diagnosed in the Great Plains. This 

oversight is unfortunate as many of the problems submitted as samples to diagnostic 

clinics are due, at least in part, to environmental stresses. Because no signs are associated 

with these disorders, identification of the specific stresses can be difficult with only a leaf 

or twig sample. An accurate diagnosis of environmental stresses requires knowledge of 

the normal appearance of the species and cultivar, its growing requirements, site conditions, and plant history. Obtaining this information may require a site visit and interview 

with the tree’s owner.

Environmental stresses are sometimes defined as only weather-related stresses; 

extremes in temperature or precipitation, or snow, ice, and wind are considered separate 

from soil-related stresses and those imposed by human activity. However, most diagnostic guides cover all these as environmental stresses but within three broad categories:

•  Human activity-related: air pollution, deicing and dust suppressant road salts, 

herbicides, improper planting, mechanical operations.

•  Soils-related: nutrient availability, soil moisture.

•  Weather-related: ice, precipitation, snow, temperature, wind.

This chapter will cover the most common environmental stresses in the Great Plains. 

Two of these stresses, nutrient deficiencies resulting from alkaline soils (chlorosis) and 

herbicide, are covered in more detail in other chapters as they are very common tree 

stresses in the Great Plains. Other environmental stresses such as ice, snow, and wind 

loading, and other mechanical stresses such as mower or string trimmer damage, are 

omitted from this chapter as they are fairly simple to identify in the field. Environmental 

stresses related to air pollution are also omitted. High ozone concentrations in the lower 

atmosphere are one of the most common air pollution concerns for trees and forests, but 

ozone injury to trees in the Great Plains is rare.

Hosts and Distribution

Environmental stresses are ubiquitous; they are not limited to specific hosts or distribution in the Great Plains. One key field diagnostic clue for determining the presence of 

environmental stresses is that they occur across tree species, genera, and families. It is 

common to find trees of differing genera expressing similar symptoms from exposure to 

the same environmental stress. This symptom pattern is rare with biotic stresses, which 

are generally limited to a specific host genus or species. 

However, different genera and species do vary in their tolerance to specific environmental stresses. For example, the susceptibility to damage from road deicing salts varies 

among genera; ashes (Fraxinus spp.) are generally tolerant and arborvitaes (Thuja spp.) 

are very sensitive to exposure. Tolerance levels can vary considerably even within a genus. Eastern white pine (Pinus strobus), for instance, is very sensitive to exposure to road 

deicing salts, whereas Austrian pine (P. nigra) is tolerant. This same variation is found for 

many other environmental stresses, so knowledge of a specific species’ tolerance to the 

suspected environmental stress is critical to the diagnostic process.

Environmental stresses are common throughout the Great Plains, but stresses related 

to soil and human activity are usually limited to specific sites. Flooding injury, for 

example, occurs in riparian areas; damage from road deicing salt is most apparent in 

trees lining roads. These symptom patterns can also be important clues for determining 

the presence of environmental stresses. Biotic stresses, such as infectious diseases, often 

spread randomly on a site depending on the tolerance or resistance of individual host 

trees to a specific pathogen. Environmental stresses usually follow regular patterns, such 

as the damage being most apparent just above the snow line on a marginally hardy shrub 
USDA Forest Service RMRS-GTR-335. 2016. 5

Figure 1-1—A common dieback pattern associated with 

abiotic stresses. The definite line to the dieback here was the 

snow depth on the privet hedge that protected the lower buds 

from winter injury (John Ball, South Dakota State University, 

used with permission).

(fig. 1-1). It is much easier to identify environmental stresses in the field, rather than from 

samples, because a field visit allows symptom patterns to be observed and information to 

be gathered on the plant’s history. 

Symptoms

No signs are associated with environmental stresses. Instead, symptoms, which are 

physical changes in the tree’s appearance, are used in the field to determine the presence 

and role of these stresses. But symptoms are not as diagnostic as signs; very similar 

symptoms may be associated with different environmental stresses (table 1-1). Marginal 

necrosis (scorch)—blackened or brown tissue along the edge of the leaf blade—may be 

the result of a late spring frost or a summer drought. Symptoms associated with environmental stresses may also mimic those expressed by injury from insects or pathogens, 

thereby complicating diagnosis of their role. For example, a stunted pine shoot may 

be a result of drought, road 

deicing salts, pathogens, or 

borers. Another difficulty in the 

diagnostic process is that environmental stresses may not be 

limited to a single stress agent, 

but may occur as multiple 

stresses, such as drought and 

saline soils. The combination 

of two or more environmental 

stresses may result in a symptom pattern different from that 

of a single stress acting alone.

Determination of the stress 

or stresses responsible for tree 

damage may require further 

investigation in a laboratory. 

Although the symptom pattern 

on roadside trees may indicate 

use of road deicing salts, tissue samples may be necessary to determine the presence or 

absence of salts and whether their concentrations were sufficient to cause the damage 

observed. This step may also be required when associating symptoms with nutrient deficiencies or herbicide drift. 

Table 1-1—Common environmental stress agents and their symptoms

Foliage symptoms Canopy symptoms

Premature Premature Reduced

Environmental autumn foliage Small Adventitious shoot Split

stress agent Chlorosis color drop Scorch foliage Wilting shoots Dieback growth trunk

Alkaline soils— 

iron/manganese 

deficiencies x x x x x

Drought x x x x x x

Flooding x x x x x x x

Freeze x x x x x

Frost x x x x x x

Planting errors x x x x x x x

Poorly drained soils x x x x x

Road deicing/dust 

suppressant salts x x x x x

Saline soils x x x x x

Stem-girdling roots x x x x x x
6 USDA Forest Service RMRS-GTR-335. 2016.

Cause and Damage

Alkalinity and iron or manganese availability 

Chlorosis is covered in more detail in chapter 5 of this publication, but the symptoms 

are so prevalent in Great Plains trees that it is worth mentioning in this chapter as well. 

Most tree species perform best in soils that are slightly acid (pH 6.1 to 6.5). Soil pH 

greater or less than this range can result in excessive or reduced availability of specific 

soil nutrients. Slightly alkaline soils, those with a pH between 7.4 and 7.8, are common in 

the Great Plains. These soils may lack sufficient available (soluble) iron or manganese to 

support certain tree species. 

Two of the species most sensitive to reduced availability of micronutrients are pin oak 

(Quercus palustris) and red maple (Acer rubrum). When planted on slightly alkaline soils 

or even neutral soils, these trees express symptoms of interveinal chlorosis, which is yellowing of the leaf tissue surrounding the veins (fig. 1-2). There is also a gradient of these 

symptoms along a branch, with the symptoms more severe on the newest foliage. These 

micronutrients are immobile, so they cannot be transferred from older to newer foliage 

tissue. 

Most tree species can tolerate slightly alkaline soils and there are a few tree species 

such as black walnut (Julgans nigra) and bur oak (Q. macrocarpa) that may not show 

any symptoms of chlorosis until the soil pH exceeds 8.0. Although chlorosis is most often 

linked to alkaline soils, it can also appear in trees on saline or poorly drained soils. There 

are also tree cultivars grown in the Great Plains that have yellow or golden ornamental 

foliage, which may be confused with chlorosis. These cultivars include ‘Princeton Gold’ 

Norway maple (A. platanoides ‘Princeton Gold’), Golden Shadow® pagoda dogwood 

(Cornus alternifolia ‘W. Stackman’), ‘Taylor’s Sunburst’ pine (P. contorta ‘Taylor’s 

Sunburst’), and golden red oak (Q. rubra ‘Aurea’).

Drought/heat

Excessive heat and water deficit stresses usually occur in combination, so the two are 

generally discussed together with water deficits considered the more serious stress of the 

two. Deciduous trees affected by water deficits may exhibit foliage that droops, curls, and 

become scorched (fig. 1-3). The scorching will usually move deeper into the leaf tissue 

between the veins. Severely injured foliage will discolor, often turning yellow to brown, 

and abscise prematurely. Conifer needles may turn brown, droop slightly near the base, 

and shed prematurely. Pines affected by long-term drought may retain only their one- and 

two-year-old needles.

Shoot extension may be reduced either during the year of the drought or the next year 

depending upon the season the drought occurred and whether the affected species has 

preformed shoot growth (the environmental conditions one year determine the extent of 

shoot growth the next year) or sustained shoot growth (the conditions during the present year determine the growth that same year). Trees such as pine and spruce (Picea 

spp.) that exhibit preformed growth will produce abnormally shortened needles and 

shoots the year following a drought (fig. 1-4). This “bottlebrush” growth pattern is often 

Figure 1-2—Chlorotic leaf from a swamp white 

oak (Q. bicolor) (John Ball, South Dakota State 

University, used with permission).

Figure 1-3—Browning and wilting oak leaves 

affected by drought (John Ball, South Dakota 

State University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 7

accompanied by excessive production of cones, both of which are very common symptoms of water deficits from the previous growing season. Trees such as birch (Betula 

spp.) that have sustained growth will show reduced shoot growth during the summer 

when the drought occurred. However, symptoms of late-summer and fall drought injury 

may not appear until the next year. Late-season drought may also increase the potential 

for winter freeze injury.

Flooding

Flooding is a frequent occurrence along the many streams and rivers in the Great 

Plains. There are many factors to consider when determining the impact of flooding on a 

tree. Early spring flooding that occurs while the trees are dormant imposes less stress than 

flooding that occurs during the growing season. Colder and fast-moving water causes less 

injury than warm, stagnant water. Flooding depth is also a factor, with water covering 

the trunk or canopy more injurious than a flood that only saturates the soil. Season and 

duration of flooding are factors as well. Many trees are able to tolerate spring flooding 

for several days. Most tree species can survive flooding for several weeks, but few can 

survive for time periods longer than two months. The list of species that can survive long 

flooding periods includes boxelder (A. negundo), green ash (F. pennsylvanica), plains cottonwood (P. deltoides ssp. monilifera) and willow (Salix spp.). Flooding symptoms often 

appear first as premature fall foliage color followed by wilting (fig. 1-5). Dieback and 

decline of the canopy may follow, depending upon the species’ tolerance to flooding and 

the depth, duration, and season of the flood. 

Frost

Late frost may occur while a tree is breaking bud and the tender shoots and leaves 

are beginning to expand. This tissue is vulnerable to cold temperatures and if exposed to 

freezing temperatures, it can be injured or killed. Late spring frosts result in wilted and 

scorched foliage on deciduous trees (fig. 1-6) and curled and distorted needles on conifers. Stems of young deciduous trees may develop vertical splits.

Figure 1-4— “Bottlebrush” appearance 

of blue spruce (Picea pungens) shoots, 

an indication of past stresses (John Ball, 

South Dakota State University, used with 

permission).

Figure 1-5—Premature fall foliage color, a 

common symptom of flooding injury (John 

Ball, South Dakota State University, used with 

permission).

Figure 1-6—Wilting of newly expanding foliage, 

a common symptom associated with late frosts 

(John Ball, South Dakota State University, used 

with permission).
8 USDA Forest Service RMRS-GTR-335. 2016.

Planting errors

Improper planting is another common environmental stress agent. Errors at planting 

cannot be easily corrected later and may result in long-term growth reduction, decline, 

and premature death. The most common long-term planting stress is from planting too 

deep. Placing soil around the trunk and covering the trunk/root interface can result in a 

slow decline. A tree that was planted too deep may persist for years, even decades, but 

the shoot growth is usually much less than normal and the leaves may be small. The most 

common clue that the tree has been planted too deep is that the stem comes straight out 

of the ground without the wider taper that normally occurs at the flare. Most ornamental 

trees are budded, so the bulge or crook associated with the budding onto a rootstock 

should still be above ground on young trees (fig. 1-7). 

Poorly drained soils

High precipitation combined with clay soils or a perched water table can result in excessive soil moisture, which contributes to root decline. Trees growing on poorly drained 

soils may appear stunted and the foliage may become chlorotic (fig. 1-8). The foliage 

may also turn red or yellow by late summer. Some tree species are less adapted to this 

environment than others. Spruces are poorly suited to such sites, whereas willows may 

thrive. 

Road deicing and dust suppressant salt applications

Deicing and dust suppressant road treatments involve applications of liquid or solid 

forms of magnesium chloride or sodium chloride. The chlorine ion is considered to be the 

most injurious to trees. Chloride can be deposited on the buds and foliage and absorbed 

into the tissue or enter the tree through the root system from salt runoff. Symptoms of 

chloride toxicity on conifers typically begin as browning foliage, starting at the needle 

tips, and on the side of the tree facing the road (fig. 1-9). The injured foliage will shed 

prematurely so that perhaps only the one- and two-year-old needles persist. Deciduous 

trees will have buds fail to open, resulting in branch dieback. Witches’-brooms, which are 

clusters of adventitious shoots, usually occur on the side of the canopy facing the road. 

Some tree species are more sensitive to these salts: little-leaf lindens (Tilia cordata) may 

be affected, yet nearby honeylocust (Gleditsia triacanthos) do not show any symptoms.

Saline soils

Some regions of the Great Plains have soils that are naturally high in salts or become 

saline from soluble salts in irrigation water. High salt concentrations can disrupt the 

movement of water from the soil into the roots. High soil salt concentrations can also 

change soil structure. The salts can bind to clay particles, reducing pore spaces and 

thereby potentially compromising gas exchange and water drainage. Damage from saline 

soils most often occurs in seedlings rather than in mature trees. Tree seedlings in saline 

Figure 1-7—A tree that was planted too deep. 

The slight bulge at the base of this budded tree 

should have been above the soil surface (John 

Ball South Dakota State University, used with 

permission).

Figure 1-8—Declining spruce in poorly drained 

soils (John Ball South Dakota State University, 

used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 9

soils may exhibit foliage scorch or wilting. 

The trees may also become stunted or die 

if soil salt concentrations are too high. The 

soil salt concentration can be determined 

only by soil testing, and soils with a 

conductivity greater than 8 deciSiemens/

meter (dS/m) measured as a saturated paste 

are harmful to all but two tree species, 

Russian-olive (Elaeagnus angustifolia) and 

mugo pine (P. mugo). Many tree seedlings 

can become established and grow on 

slightly saline soils, that is, those with a 

conductivity near 4 dS/m.

Stem-girdling roots

Associated with burying the trunk in 

the soil at planting is the development of 

stem-girdling roots. The development of 

a stem-girdling root requires a portion of 

the stem be below grade, which allows a 

root to rise to constrict the stem (fig. 1-10). 

Stem-girdling roots are more common 

in species that generate roots that grow 

tangential to the trunk following harvest 

or root pruning. Maple, ash, and linden are the three most common genera associated 

with this disorder. Stem-girdling roots may also occur with root-bound container stock 

if the circling roots are not shaved off at planting. Stem-girdling roots can reduce tree 

stability because of the distorted root system. The girdling can also restrict photosynthate 

movement to the roots, and water and 

nutrient movement to the foliage. The 

most common symptoms associated 

with stem-girdling roots are leaf scorch, 

abnormally small foliage, premature fall 

foliage color, premature foliage drop, 

dieback, and death. The stem may also 

appear flattened along one side.

Winter injury

Planting a species farther north than 

its native range may result in cold winter 

temperatures killing the tree or, as is 

often the case with marginally hardy tree 

species, branch dieback, which provides 

an entry wound for pathogens. Thornless 

honeylocust (G. triacanthos var. inermis) 

cultivars that originate from trees native to milder climates may be injured by the cold 

winter temperatures in the Great Plains and become more susceptible to canker diseases. 

The different parts of the tree—buds, sapwood, roots—acclimate to different midwinter minimal temperatures, so cold injury might be limited to the loss of specific tissue. 

The sapwood is less cold tolerant than the vegetative buds. Cold injury to the sapwood 

may not be noticed until the spring, when the expanding foliage on the affected shoot 

wilts and dies. A brown streak is often present in the sapwood of these injured trees. 

Roots are the more cold-sensitive part of a tree and the woody roots for many tree 

species cannot tolerate temperatures below 15 °F. Cold injury to roots can occur during 

winter on sites with bare soils and inadequate snow cover. Trees with extensive cold injury to the woody roots will often partially leaf out in spring before wilting. This injury can 

occur on tree cultivars that are generally considered hardy to the region if the rootstock is 

Figure 1-9—A ponderosa pine (Pinus ponderosa) exhibiting symptoms of injury from road 

de-icing salts (John Ball South Dakota State 

University, used with permission).

Figure 1-10—A stem-girdling root appearing on 

a tree (John Ball, South Dakota State University, 

used with permission).
10 USDA Forest Service RMRS-GTR-335. 2016.

from a more southern seed source or even from a cold-sensitive species. Winter injury has 

been observed in the Great Plains when State Street® maple (A. miyabei ‘Morton’), which 

is hardy to much of the region, is budded onto hedge maple (A. campestre) rootstock, a 

tree hardy only to the southern Great Plains.

Much of the winter injury in the Great Plains does not occur in midwinter but occurs 

during late fall and early spring as trees are acclimating to or de-acclimating from freezing temperatures. The timing and rate of acclimating to cold winter temperatures differ 

among species and may even differ among seed sources within a species. As the photoperiod shortens in late summer and early fall, trees begin the process of acclimation to the 

subfreezing conditions of winter. This process accelerates with exposure to freezing night 

temperatures. If the fall is abnormally warm with very few freeze/thaw cycles before cold 

winter temperatures begin, the trees may not be fully acclimated. Injury can also occur 

in late winter and early spring when trees are deacclimating and preparing to resume 

growth. If a tree has fulfilled its chilling requirement—exposure to a minimum period 

of cold temperatures—and the weather turns unseasonably warm then back to morenormal cold temperatures, freeze injury may result. Because of the continental climate 

of the Great Plains, this region experiences more unseasonable temperature fluctuations 

between fall and spring than areas along the east or west coast, where oceans help in 

moderating temperatures. 

A type of winter injury common to evergreens is winter desiccation injury, commonly 

referred to as winter burn. This injury occurs during sunny and windy winter days where 

temperatures are high enough that water can be lost to transpiration. This water cannot be 

replaced as the soil and trunk are still 

frozen, so the foliage desiccates. The 

injured foliage turns red or brown (fig.

1-11), though this color change may 

not occur until spring as the evergreen 

begins to resume growth.

Management

Environmental stresses are common, 

but often overlooked, causal agents 

when diagnosing tree problems. They 

are easy to miss as environmental 

stresses do not produce signs, only 

symptoms, and these symptoms can 

also be associated with multiple abiotic 

and biotic agents. Identification of 

environmental stresses requires not only 

knowledge of the symptoms associated 

with a particular stress but also site information and history. For example, are the soils 

alkaline or saline, or was there a late frost the past spring? Determining environmental 

stresses is difficult to do from a sample or a picture attached to an email and often requires site information and a history of the plant’s planting and care.

Another reason that environmental stresses may be overlooked is their underlying role 

in a tree’s decline. Although environmental stresses can be primary factors in tree mortality, they more often serve as predisposing factors in tree decline. The gradual decline of 

a tree can be described as a mortality spiral as the tree’s susceptibility and sensitivity to 

abiotic and biotic agents increase as the tree declines. The spiral begins with predisposing 

factors, followed by inciting and finally contributing factors. Predisposing factors are 

long-term stresses that reduce the tree’s health on a particular site and increase susceptibility to pests. The most common predisposing factors in tree decline are environmental 

stresses such as poorly drained soils and planting errors. 

Trees may endure predisposing stresses for many years until an inciting factor occurs. 

Incitants are short-duration stress agents that further compromise the tree’s health and its 

capability to defend itself from pests. Common incitants are late frosts and drought, or 

defoliation by leaf-feeding insects or pathogens. These inciting factors add to the stresses 

Figure 1-11—Browning of concolor fir needles 

from winter burn (John Ball, South Dakota State 

University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 11

imposed by the predisposing factors and the combined effect will be reduced capability of 

the tree to defend against secondary pests.

Secondary pests, referred to as contributing factors in the decline spiral, are generally 

biotic; common examples are bark beetles, root diseases, and canker diseases. Detection 

of these organisms in a tree often coincides with symptoms of extensive canopy dieback. 

The observer may associate the appearance of dead branches with the presence of borers 

or cankers. These organisms often are assumed to be the sole cause of the decline and the 

predisposing factors and incitants ignored. 

Making this assumption often results in managing only the contributing factors. 

Merely treating the contributing factors may not arrest the tree’s decline. Tree managers 

should instead address or at least acknowledge the underlying stresses, both predisposing 

and inciting, and try to understand their role in the tree’s decline. 

Once the environmental stresses are identified, there is often little that can be done 

to alleviate their effects. A marginally cold-hardy tree subjected to winter injury will be 

injured every winter. Predisposing factors are not usually correctable or alleviated by any 

treatment. Environmental inciting factors, though not correctable, may be mitigated by 

tree care practices such as watering and mulching to reduce their impact.

Selected References

Ball, J. 2013. Trees! Fruit, nut, ornamental, shade, and windbreak trees for the Northern 

Plains. Brookings, SD: Igrow Publishing. 494 p.

Clark, J.R.; Matheny, N. 1991. Management of mature trees. Journal of Arboriculture. 17: 

173–184.

Luley, C.J.; Ali, A.D. 2009. Pest management in the landscape: an introduction. Naples, 

NY: Urban Forestry LLC. 89 p.

Manion, P.D. 1991. Tree disease concepts. 2nd ed. Englewood Cliff, NJ: Prentice-Hall, 

Inc. 416 p.

Mittler, R. 2006. Abiotic stress; the field environment and stress combination. Trends in 

Plant Science. 11: 15–19.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Schoeneweiss, D. 1981. The role of environmental stress in diseases of woody plants. 

Plant Disease. 65: 308–314.

Vollenweider, P.; Gunthardt-Goerg, M.S. 2005. Diagnosis of abiotic and biotic stress 

factors using the visible symptoms in foliage. Environmental Pollution. 137: 455–465.
12 USDA Forest Service RMRS-GTR-335. 2016.

2. Anthracnose Diseases of Broadleaf Trees

Scott C. Redlin; revised from Robert W. Stack and 

Kenneth E. Conway (Riffle and Peterson 1986)

Many trees are affected by diseases caused by fungi with asexual stages that produce 

asexual spores (conidia) in acervular conidiomata. Diseases caused by these fungi 

are commonly called anthracnose and are usually connected to Ascomycete fungi 

(Diaporthales: Gnomoniaceae1). The site of infection, symptoms and signs, and severity 

of the anthracnose disease differ from species to species and often vary from year to year. 

The only symptom commonly found on maples (Acer spp.) and elms (Ulmus spp.) may 

be leaf spot or leaf blight (figs. 2-1 and 2-2). In other species, such as sycamore (Platanus

spp.) and white oaks (Quercus spp. such as Q. macrocarpa), the entire range of anthracnose symptoms may occur, including bud, twig, and shoot blights, leaf spots, and stem 

cankers. Several of the causal fungi are now considered latently infecting or endophytic. 

Fungi on host trees have asexual stages in several genera including Asteroma, Discula, 

and Marssoniella. Some species have two or more anamorphic stages (synanamorphs). 

Numerous names for the anamorphs of Apiognomonia errabunda, which occurs on basswood (Tilia spp.), oak, and other hosts have been proposed, but few are nomenclaturally 

acceptable.

Hosts and Distribution

Anthracnose diseases caused by Gnomoniaceae are widespread throughout the native 

and planted ranges of their respective hosts. The pathogens and their hosts growing in the 

Great Plains are summarized in table 2-1. Each fungal species is specific for a particular 

host genus, with little or no cross pathogenicity as far as is known. The taxonomy of the 

causal fungi of anthracnose diseases has been revised. For each host, the symptoms may 

differ based on the part of the tree attacked (table 2-1). 

Symptoms and Signs

Leaf infections early in the season result in blotching, distortion, and large necrotic 

areas. These symptoms are typical of ash (Fraxinus spp.), oak, and sycamore anthracnose 

(fig. 2-3 through fig. 2-6). If symptoms occur early in the season, they can be confused 

with frost damage. Later leaf infections result in discrete small to large spots, often 

1 The fungi in the order Diaporthales causing anthracnose symptoms discussed here have been 

proposed to be reclassified to include the families Sydowiellaceae and Valsaceae in addition to 

Gnomoniaceae.

Figure 2-2—Symptoms of maple anthracnose caused by Aureobasidium apocryptum 

(Paul Bachi, University of Kentucky, used with 

permission).

Figure 2-1—Black leaf spot of elm caused 

by Stegophora ulmea (Bruce Watt, 

University of Maine, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 13

Table 2-1—Pathogens causing anthracnose diseases of trees in the Great Plains 

Host attacked Name of pathogena Parts of tree attacked

Ash, especially green ash Plagiostoma fraxini Leaves, twigs, samaras

Syn.: Gnomoniella fraxini

Anamorph: Discula fraxinea

Basswood Apiognomonia errabunda Leaves, twigs

Anamorph: Discula sp.

Elm Stegophora ulmea Leaves 

Anamorph: Asteroma ulmea

Maple Aureobasidium apocryptum Leaves

Colletotrichum gloeosporioides

Discula campestris

Discula spp.

Oak, especially white oaks Apiognomonia quercinab Leaves, twigs, shoots, 

Anamorph: Discula quercina buds

American sycamore; Apiognomonia veneta Leaves, twigs, shoots,

London plane-tree Anamorph: Discula platani buds 

Walnut Ophiognomonia leptostyla Leaves, twigs, nuts

Anamorph: Marssoniella

juglandis (syn.: Marssonina

juglandis)

a The anamorph names of most of the anthracnose fungi require further study for accurate taxonomic 

placement. They are provided in table 2-1 because the anamorph, commonly observed during spring 

and summer, is the basis of disease diagnosis. 

b Amphiporthe leiphaemia is proposed as the most recent nomenclature for this pathogen.

Figure 2-3—Leaf distortion and necrosis of 

green ash caused by Plagiostoma fraxini

(Michael Kangas, North Dakota Forest 

Service, used with permission).

Figure 2-4—Acervuli of Apiognomonia quercina

on midvein of oak (Bruce Watt, University of 

Maine, used with permission).

surrounded by a dark ring or a chlorotic halo. Such spots are the principal symptoms in 

basswood, elm, and maple (figs. 2-1 and 2-2). Severe leaf spot infection may cause extensive defoliation, especially in ash, sycamore, and walnut (Juglans spp.). 

On sycamore, four phases of anthracnose 

can be distinguished: twig blight, bud blight, 

shoot blight, and leaf blight. Leaf blight is 

already described above. In the twig blight 

phase, small, 1- or 2-year-old twigs are girdled 

and killed (fig. 2-5). Later, larger twigs and 

branches may be cankered and killed. In the 
14 USDA Forest Service RMRS-GTR-335. 2016.

Figure 2-6—Leaf necrosis of sycamore 

caused by Apiognomonia veneta (American 

Phytopathological Society, used with permission). 

Figure 2-5—Leaf necrosis and twig dieback 

of sycamore caused by Apiognomonia veneta 

(Joseph O’Brien, U.S. Forest Service).

bud blight phase, buds are penetrated by the fungus and killed before they expand. In 

the shoot blight phase, new young shoots and expanding leaves are suddenly killed. In 

sycamore (and probably other hosts), severe shoot blight depends on temperature, with 

infection greatest when temperatures are 50 to 59 °F. In walnut, optimal infection temperature is 70 °F.

Disease Cycle

The life cycles of these anthracnose fungi are similar in that they require water from 

rain, dew, or fog to infect a tree. However, most anthracnose fungi do not incite all four 

phases of symptoms, and Apiognomonia veneta does not represent the typical life cycle in 

the Midwest. The fungi survive the winter primarily on fallen infected leaves and sometimes in infected twigs and branches. With ash anthracnose, petioles colonized by the 

powdery mildew fungus, Phyllactinia fraxini, often persist through winter and provide a 

major source of anthracnose inoculum close to the buds, from which susceptible shoots 

emerge in spring. In most situations there is abundant inoculum for infection if conditions 

are favorable. Asexual spores (conidia) are the most important stage for dissemination 

and infection by anthracnose fungi. In several species, the sexual spores (ascospores) 

may also be produced and serve as primary inoculum; however, their relative importance 

compared to the conidial stage is undetermined.

Spores are disseminated by wind and splashing rain to buds, shoots, and expanding 

leaves, where infection begins if conditions are favorable. In some species, infections develop during favorable periods in summer, giving rise to late-season spots or leaf blight.

Damage

Anthracnose fungi can cause defoliation and contribute to branch dieback, which 

disfigures the tree. If anthracnose is severe for several seasons, the tree may be weakened 

and start to decline, or may become susceptible to other diseases or insect pests. Trees 

subject to other stresses such as root restriction, drought, or heavy scale infestation, are 

much less tolerant to anthracnose, and may show decreased vigor after only a single season of severe anthracnose defoliation.

Management

Because individual trees may vary in susceptibility to anthracnose, there is potential 

for selection of resistant clones or seed sources of highly resistant trees. The true London 

plane-tree (an interspecific hybrid, Platanus x acerifolia) is much less susceptible than 

the native American sycamore (Platanus occidentalis). Oaks in the white oak group are 

much more susceptible to anthracnose than those in the red oak group (for example, 

Q. palustris). 

Several cultural practices can help manage anthracnose diseases and include sanitation and pruning to remove infected twigs and branches during the dormant period. In 

walnut anthracnose, removing some branches by pruning increases air movement through 

crowns, facilitates faster drying of leaves after rain, and reduces infection of leaves. 
USDA Forest Service RMRS-GTR-335. 2016. 15

Application of nitrogenous fertilizers promotes tree vigor and could result in less-severe 

disease. 

Fungicide sprays have been used in attempts to control anthracnose on boulevard and 

residential plantings but are rarely warranted because anthracnose diseases do not occur 

every year and seldom seriously affect tree health. Specific protectant fungicides are 

registered for certain tree species. Follow label instructions for times of application and 

dosage rates. Specific recommendations on type of spray and timing depend on locality. 

Consult your local extension plant pathologist or forester for current recommendations 

for your area. Several systemic fungicides have been registered for tree injection to control anthracnose fungi.

Selected References

Hepting, G.H. 1971. Diseases of forest and shade trees of the United States and Canada. 

Agric. Handb. 386. Washington, DC: U.S. Department of Agriculture. 658 p.

Himelick, E.B.; Neely, D. 1988. Systemic chemical control of sycamore anthracnose. 

Journal of Arboriculture. 14: 137–141.

McGranahan G.; Smalley, E.B. 1984. Conidial morphology, axenic growth and 

sporulation of Stegophora ulmea. Phytopathology. 74: 1300–1303.

Mejía, L.C.; Castlebury, L.A.; Rossman, A.Y.; Sogonov, M.V.; White, J.F., Jr. 2011. A 

systematic account of the genus Plagiostoma (Gnomoniaceae, Diaporthales) based on 

morphology, host-associations, and a four-gene phylogeny. Studies in Mycology. 68: 

211–235. 

Monod, M. 1983. Monographie taxonomique des Gnomoniaceae. Beihefte zur Sydowia. 

Annales Mycologici, Series 2. 9: 1–315.

Redlin, S.C.; Stack, R.W. 1988. Gnomoniella fraxini sp. nov., teleomorph of the ash 

anthracnose fungus and its connection to Discula fraxinea comb. nov. Mycotaxon. 32: 

175–198. 

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p. 

Sogonov, M.V.; Castlebury, L.A.; Rossman, A.Y.; Mejía, L.C.; White, J.F. 2008. Leafinhabiting genera of the Gnomoniaceae, Diaporthales. Studies in Mycology. 62: 1–77.

Sogonov, M.V.; Castlebury, L.A.; Rossman, A.Y.; White, J.F. 2007. The type species of 

Apiognomonia, A. veneta, with its Discula anamorph is distinct from A. errabunda. 

Mycological Research. 111: 693–709. 

Sutton, B.C. 1980. The Coelomycetes: Fungi Imperfecti with pycnidia, acervuli and 

stromata. Kew, Surrey, UK: Commonwealth Mycological Institute. 696 p. 
16 USDA Forest Service RMRS-GTR-335. 2016.

3. Bur Oak Blight

Thomas C. Harrington and Douglas L. McNew

Many species of Tubakia are capable of causing late-season leaf spots and necrosis of 

leaf veins on oak (Quercus spp.), chestnut (Castanea spp.), and other hardwood species 

in Europe, Asia, and North America. But the recently described fungus T. iowensis is 

also capable of causing petiole necrosis and death of whole leaves on bur oak (Q. macrocarpa), sometimes killing nearly every leaf on a susceptible tree. Damage to bur oak was 

first noted in the mid-1990s in the northern Great Plains and the Upper Midwest, and the 

incidence appears to be increasing in this region, perhaps due to more abundant spring 

rainfall.

Hosts and Distribution

Substantial necrosis of leaf veins and petioles by T. iowensis occurs only on bur oak. 

Bur oak blight is most common in Iowa and Minnesota, but the disease has been noted in 

western and southern Wisconsin, northern Illinois, northeast Kansas, eastern Nebraska, 

and eastern South Dakota, with isolated groups of affected trees in counties of Illinois and 

Missouri that border Iowa. This disease distribution roughly coincides with the limited 

distribution of a small-acorned, fire-adapted variety of bur oak, Q. macrocarpa var. oliviformis. Bur oak blight is most severe in relic savanna groves on upland sites (fig. 3-1). 

On bottomland sites, few trees show severe symptoms, and the disease has not been 

recorded where the large-acorned Q. macrocarpa var. macrocarpa is common. The disease does not seem to be present in eastern 

North Dakota, where Q. macrocarpa var. 

depressa predominates on sites with sandy 

soils. Ornamental plantings of bur oak trees 

with large acorns are not seriously affected, 

but planted bur oak with small acorns may 

have severe symptoms (fig. 3-2).

Symptoms and Signs

The first symptoms usually appear in June 

in the form of elongated, purple-brown 

lesions on veins of the leaf undersurface, 

sometimes with numerous small, reddishbrown necrotic spots on the leaf blade 

between veins (fig. 3-3). Sporulation is 

not evident until there is more substantial 

necrosis of the leaf veins in late June and 

throughout summer (fig. 3-4), when leaf 

symptoms could be mistaken for anthracnose caused by Amphiporthe leiphaemia 

(formerly Discula quercina). Fruiting structures that bear asexual spores (conidia) of 

T. iowensis appear on or near the necrotic 

veins. On the upper leaf surface, masses of 

conidia are typically covered with a shield 

of red-brown, radiating, setal-like structures (fig. 3-5). Unshielded conidial masses 

form on the veins on the underside of the 

leaves. The necrosis may expand down to 

the base of the leaves, and the leaves are 

shed. Substantial defoliation may be seen 

during wet summers (fig. 3-6). Veinal necrosis and fruiting structures with radiating 

shields are evident on the fallen leaves. 

Figure 3-1—Two mature severely blighted 

bur oak trees (left). Other bur oak trees in the 

grove appear healthy, presumably because of 

resistance (Thomas C. Harrington, Iowa State 

University, used with permission).

Figure 3-2—A planted bur oak tree with small 

acorns, showing substantial leaf symptoms in 

the lower crown (Thomas C. Harrington, Iowa 

State University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 17

Figure 3-3—Initial symptoms of bur oak blight 

in June: purple-brown lesions on the veins on 

the underside of leaves (Thomas C. Harrington, 

Iowa State University, used with permission).

Figure 3-4—Midsummer symptoms of bur oak 

blight: veinal necrosis, with the fungus producing 

shielded masses of conidia along the necrotic 

veins on the upper surface and naked conidial 

masses on the veins of the lower leaf surface 

(Thomas C. Harrington, Iowa State University, 

used with permission).

Figure 3-5—Top view of a fruiting structure 

of Tubakia iowensis from the upper leaf 

surface, showing the shield of radiating, pigmented hyphae and conidia produced under 

the shield (Douglas L. McNew, Iowa State 

University, used with permission).

Figure 3-6—Bur oak blight-infected trees with 

thin crowns. In wet summers, infected trees may 

shed most of their symptomatic leaves (Thomas 

C. Harrington, Iowa State University, used with 

permission).

Coincidental with veinal necrosis and 

defoliation, necrosis on the base of the 

petiole and death of whole leaves appear 

in late July and into August (fig. 3-7). 

Such leaves may remain hanging on the 

tree, and black pustules begin to form 

in late summer on the necrotic petiole 

tissue. However, these pustules typically 

do not mature and produce conidia until 

the next spring, at the time of leaf emergence (fig. 3-8). Healthy bur oak branches typically shed all of their leaves heading into winter. 

The fruiting structures and conidia of T. iowensis (fig. 3-5) are difficult to distinguish 

from those of other Tubakia species. However, the shielded fruiting structures along the 

veins on the upper leaf surface and naked conidial masses on the veins of the lower leaf 

surface are typical for T. iownesis. Only T. iowensis forms black, crustose fruiting structures on the lower 0.4 inch of overwintering petioles of bur oak (fig. 3-7), but other fungi, 

such as Botryosphaeria species, may form similar black structures on midveins, petioles, 

and twigs. A twig blight associated with B. corticola may also leave dead leaves hanging 

on branches throughout the winter, but in this case the dead leaves are clustered on dead 

branch tips instead of scattered among the twigs. The conidia of Botryosphaeria species 

are much larger than those of T. iowensis.
18 USDA Forest Service RMRS-GTR-335. 2016.

Figure 3-7—Leaves endophytically infected by Tubakia iowensis. Necrosis of petioles may eventually be shown, leading 

to dead leaves that remain firmly attached to branches 

until the next spring, when conidia are produced from black 

pustules on the petioles (Thomas C. Harrington, Iowa State 

University, used with permission).

Figure 3-8—Black, crustose pustules of Tubakia iowensis on the petioles of overwintering leaves. Pustules 

produce rain-splashed conidia at the time of shoot 

emergence in spring (Douglas L. McNew, Iowa State 

University, used with permission).

Dead leaves and leaves with 

veinal necrosis are typically most 

evident in the lower crown (fig. 

3-2). Trees severely defoliated 

in consecutive years may have 

thin crowns, and twig death and 

branch dieback are sometimes 

seen. However, the branch 

dieback is most likely associated 

with the two-lined chestnut borer, Agrilus bilineatus. Branch dieback associated with the 

two-lined chestnut borer is commonly found on trees affected by drought, root rots, and 

other primary stresses, so branch dieback is not diagnostic for bur oak blight. It is likely 

that some trees with bur oak blight have been misdiagnosed as oak wilt, which may also 

result in dead foliage and branch dieback. 

Disease Cycle

Black, crustose pustules formed on last season’s petioles provide the initial inoculum 

in spring. The fungus may produce sexual spores (ascospores) in spring, but these have 

not been found and are not likely of major importance. The petiole pustules on hanging 

leaves expand and break open with wetness in spring, and the conidia are dispersed by 

rain, leading to endophytic and symptomless infections of expanding shoots and leaves 

(fig. 3-8). After wet springs, nearly every leaf on a susceptible tree is endophytically 

infected but remains symptomless for two months or more, when petiole necrosis may 

become evident. This petiole necrosis apparently prevents normal leaf abscission in the 

fall.

When mature leaves are infected during the summer, veinal necrosis and leaf death 

may be conspicuous, and the fungus produces abundant conidia on both the upper and 

lower leaf surfaces. Many leaves infected during the summer are shed, and it is not clear 

if summer-infected leaves remain attached and produce petiole pustules the next spring. 

Wet summers tend to lead to more veinal necrosis and defoliation, but dry summers 

following wet springs may lead to more petiole necrosis and many dead leaves hanging 

on the tree. Consecutive years of normal or above-normal spring rainfall seem to lead to 

buildup of the disease in individual trees. The conidia are dispersed by rainsplash, which 

might be why the lower crowns of the trees are typically more seriously affected than the 

upper crowns (figs. 3-2 and 3-6).
USDA Forest Service RMRS-GTR-335. 2016. 19

Damage

Bur oak blight has been most severe in former savanna groves. However, both the host 

and pathogen appear to be native, and even in the most severely affected groves, many of 

the trees show few or no symptoms (fig. 3-1), presumably because of resistance. Within 

the known range of the disease, fewer trees appear to be affected on bottomland sites as 

compared to upland sites, perhaps because bottomland ecotypes of bur oak tend to be 

more resistant. Most ornamental bur oak trees do not show symptoms of bur oak blight, 

but some planted trees are seriously affected (fig. 3-2). When acorns have been seen on 

planted bur oak trees with bur oak blight, the acorns have been small, typical for Q. macrocarpa var. oliviformis.

Severe symptoms in late summer appear to follow periods of high rainfall during the 

previous spring, and a recent shift in climate towards more spring rainfall is believed to 

be responsible for the relatively high incidence of the disease in Iowa and Minnesota. 

Consecutive years of high spring rainfall may lead to buildup of overwintering inoculum 

(pustules on petioles) in the crown of susceptible trees. High summer moisture may also 

lead to an abundance of secondary infections with veinal necrosis and defoliation. In 

contrast, summer drought following normal spring rains may lead to more dead leaves 

hanging on the trees.

Some trees with repeated defoliation may show twig and branch dieback, generally 

attributable to the two-lined chestnut borer. Aside from the borer damage, relatively few 

trees affected by bur oak blight appear to die from the disease. In most areas, root rots 

and stem decays are more commonly associated with branch dieback and mortality of 

mature bur oak trees.

Management

Because the initial inoculum is produced in the tree crown, sanitation of leaves on the 

ground is not believed to be an effective control. Experimental injections of fungicide 

have been found to be effective in controlling the disease in some high-value trees, but 

only trees showing moderate to severe levels of disease should be treated. Injection of 

dilute formulations of propiconazole, following directions for oak wilt control, have been 

most effective and may reduce symptoms for two or more seasons. Inject trees as soon as 

the leaves are fully formed in late May or early June, before symptoms appear. However, 

leaves and small branches of some trees may show phytotoxicity with propiconazole 

treatments, and not all treated trees show reduced disease levels. In some treated trees, 

severe symptoms did not return until the second year after treatment, and individual trees 

need not be treated more than once every three years.

For ornamental plantings in regions with bur oak blight, grow bur oak trees produced 

from large acorns as these seem to be mostly free of the disease, as are other species of 

oak. In management of existing groves, the most susceptible trees should probably be 

removed, especially if they have substantial branch dieback. In mature groves, practice 

sanitation to reduce populations of the two-lined chestnut borer. In the absence of the 

borer, the frequency of mortality from bur oak blight alone is quite low. 

Selected References

Harrington, T.C.; McNew, D.M.; Yun, H.Y. 2012. Bur oak blight, a new disease on 

Quercus macrocarpa caused by Tubakia iowensis sp. nov. Mycologia. 104: 79–92.
20 USDA Forest Service RMRS-GTR-335. 2016.

4. Cherry Leaf Spots

Gerard C. Adams, Ned Tisserat, and William R. Jacobi

Cherry leaf spot and shot-hole disease are common diseases of foliage of sweet 

(Prunus avium) and sour (P. cerasus) cherry, Nanking cherry (P. tomentosa), and 

chokecherry (P. virginiana) in the Great Plains (fig. 4-1). Trees that are susceptible to 

these diseases are planted throughout the Great Plains: fruit-bearing cherry in yards and 

orchards, ornamental Nanking cherry, and chokecherry in windbreaks and other resource 

conservation plantings.

Hosts and Distribution

Cherry leaf spot and shot-hole disease 

are caused by bacterial and fungal plant 

pathogens of several genera. All of these 

pathogens and the related diseases are 

common throughout the range of their 

hosts and are particularly important in the 

Great Plains and eastern United States.

•  Bacterial spot caused by Xanthomonas 

arboricola pv. pruni is particularly damaging in commercial stone fruit trees, as 

well as in ornamental and native species 

grown in nurseries for home garden, shelterbelt, or soil conservation plantings. This 

pathogen is becoming the more prevalent 

and damaging of the shot-hole leaf spot pathogens.

•  Bacterial canker pathogen Pseudomonas syringae pv. syringae also may cause leaf 

spots and shot-hole symptoms and cankers with gummosis; the cankers often lead to 

susceptibility to winter injury, branch breakage, and problems in replanting. 

•  Cherry leaf spot has been the common disease name for leaf spots caused by the 

fungus Blumeriella jaapii (syn. Coccomyces hiemalis and C. lutescens) on fruitbearing cherry, Nanking cherry, sandcherry (Prunus besseyi), and chokecherry. 

•  Shot-hole blight or Coryneum blight caused by the fungus Stigmina carpophila

(syn. Coryneum carpophilum, Wilsonomyces carpophilus, and Thyrostroma carpophilum) has been the more prevalent and damaging disease with shot-hole symptoms 

on cherry in the past. The typical symptoms may be confused with the other leaf 

spots and shot-hole pathogens of cherry leading to inappropriate disease management 

treatments.

•  Cercospora leaf spot caused by the fungus Mycosphaerella cerasella is generally a 

less damaging leaf spot pathogen even on commercial stone fruit trees.

Symptoms and Signs

The term “shot-hole” refers to leaf spot diseases where the necrotic tissues in small 

circular lesions on leaves drop out, leaving circular openings or holes.

Bacterial leaf spots caused by Xanthomonas arboricola pv. pruni and P. syringae pv. 

syringae are similar to those caused by B. jaapii and S. carpophila. Lesions are reddish 

brown, are angular or circular, and frequently fall out, giving infected leaves a tattered appearance (fig. 4-2). Lesions may coalesce to form large areas of necrotic tissue. Heavily 

infected leaves are chlorotic and may be shed prematurely. Pseudomonas syringae pv. 

syringae also may cause depressed black lesions on fruits, and cankers on twigs and, as 

seen in figure 4-3, branches and the main stem. Gummosis commonly occurs around 

cankers. Bacterial leaf spots can be differentiated from fungal leaf spots by crushing a 

young leaf spot in water and examining under a microscope for the diagnostic bacterial 

streaming.

Figure 4-1—Cherry leaf spot and shot-hole 

symptoms on commercial cherry (William R. 

Jacobi, Colorado State University, used with 

permission).
USDA Forest Service RMRS-GTR-335. 2016. 21

Figure 4-2—Typical bacterial leaf spot and shot-hole symptoms caused by Xanthomonas arboricola 

pv. pruni: (A) Shot-hole on leaf of sandcherry (Kevin Korus, University of Nebraska Lincoln, used 

with permission); (B) Coalesced leaf spots forming necrotic blotches on chokecherry (Ned Tisserat, 

Colorado State University, used with permission); (C) Infection of apricot (Prunus spp.) fruit (Tyre 

Proffer, Kent State University, used with permission); (D) Spring canker on twig. Spring canker on 

twigs caused by P. syringae pv. syringae would appear similar but exhibit gummosis (David Ritchie, 

North Carolina State University, used with permission).

A

C

B

D

Figure 4-3—Pseudomonas syringae pv. 

syringae bacterial canker causing copious 

gummosis and death of scaffold branch 

on residential cherry (Gerard C.Adams, 

University of Nebraska, used with 

permission).

Fungal leaf spot and shot-hole blight

caused by B. jaapii, S. carpophila, and 

M. cerasella first appear as small, chlorotic, 

or off-color (purple to reddish) circular 

spots, which later enlarge and turn brown 

and necrotic (fig. 4-4 through 4-6). Spots 

are usually 4 to 5 mm in diameter; they 

may be few in number or may become numerous enough to coalesce and form large 

irregular necrotic areas. If the leaf spots are 

caused by fungal pathogens, then during 

humid periods, whitish (for B. jaapii and 

M. cerasella) or brownish (for S. carpophila) spore masses (conidia) may be visible 

in the center of lesions. These spore masses 

are more numerous on the underside of 

leaves for B. jaapii and occur on both sides 

of the leaves for M. cerasella. To determine 

the fungal pathogen involved in cherry 

leaf spots, place cherry leaves with intact 

spots into a moist chamber for 24 hours 

and then examine the conidia formed on 

the leaf spot. The conidia of B. jaapii and 
22 USDA Forest Service RMRS-GTR-335. 2016.

Figure 4-4—Typical fungal leaf spot caused by 

Blumeriella jaapii: (A) Leaf spots; (B) Closeup 

of leaf spot in moist weather with acervuli 

oozing conidia in cirrhi; (C) microscopic view 

of long curved conidia (Paul Bachi, University 

of Kentucky, Bugwood.org, used with 

permission). 

A B

C

Figure 4-5—Typical fungal shot-hole caused by 

Stigmina carpophila: (A) Shot-hole on leaves (Whitney 

Cranshaw, Colorado State University, Bugwood.org, 

used with permission); B) Closeup of leaf necrotic spot 

before abscission (William R. Jacobi Colorado State 

University, used with permission); (C) Spots on infected 

apricot fruit (William M. Brown, Jr., Bugwood.org, used 

with permission); (D) Microscopic view of ovoid conidia 

(Jenny Glass, Washington State University, used with 

permission). 

B

D

B

C

A
USDA Forest Service RMRS-GTR-335. 2016. 23

M. cerasella are approximately 10 times longer 

than wide and readily bend into curved shapes; 

they are colorless and light brown, respectively. 

The conidia occur in a wet mass in the acervulus 

of B. jaapii and occur dry on fascicles of brown 

conidiophores in M. cerasella. The conidia of 

S. carpophila are ovoid, brown, and dry and 

have two to five thick dark-brown cross walls 

(septa) Infections increase with each episode of 

wet weather throughout the summer. Generally, 

necrotic tissues within lesions separate from the 

surrounding leaf tissues, forming characteristic shot-holes. During severe stages of disease, leaves infected by B. jaapii become highly chlorotic, and trees may appear yellow. 

Circular black lesions can also form on twigs infected by S. carpophila and later these 

enlarge and become sunken in appearance. This pathogen can infect buds, blossoms, 

and fruit (fig 4-5c). It is the infection of buds that leads to infection of twigs and canker 

formation. M. cerasella generally remains as a 1- to 2-mm-diameter reddish-brown leaf 

spot with a lighter border (fig 4-6a, b). However, spots occasionally may coalesce into 

necrotic patches. Leaf spots caused by M. cerasella can be distinguished by the presence 

of dark fascicles of conidiophores arising from a stroma (fig. 4-6).

Disease Cycle

Bacterial leaf spots of cherry 

Xanthomonas arboricola pv. pruni overwinters in buds, leaf scars, and twig cankers. 

The twig cankers in spring can be an important source of inoculum for new infections. 

P. syringae pv. syringae overwinters within cankers and also is a common epiphyte 

that occurs on the fruit, limbs, and leaves of both susceptible and nonsusceptible plants. 

Infection begins when the epiphytic bacteria increase in population owing to physiological changes in the host and favorable weather. However, the inoculum for new infection 

usually arises from the margins of cankers as bacterial ooze. 

Both bacterial pathogens are spread to leaves and cause infections in spring during 

cool, wet weather. Infection is correlated more closely with favorable weather conditions 

than with the availability of inoculum. Free-standing water (leaf wetness) is necessary for 

leaf infection. The bacteria enter stomata primarily, but also can enter lenticels, nectaries, 

hydathodes, leaf scars, and wounds. Wounds, such as those made during pruning, are 

common infection courts. Repeated infections develop throughout the growing season 

during wet periods. Heavily infected leaves fall prematurely. 

Figure 4-6—Typical fungal leaf spot and 

shot-hole caused by Mycosphaerella cerasella: (A) Leaf spots; (B) Closeup of leaf spot; 

(C) Microscopic view of long curved conidia 

(Paul Bachi, University of Kentucky, Bugwood.

org, used with permission). 

A

B

C
24 USDA Forest Service RMRS-GTR-335. 2016.

Fungal leaf spots of cherry 

Cherry leaf spot—Infection by B. jaapii begins in spring, and is caused by ascospores from fruiting bodies (apothecia) that form on previously infected, fallen leaves. 

Ascospores are wind dispersed and cause infection during April and May in the southern 

Great Plains. Leaves are especially susceptible shortly after budbreak. The summer form 

of the pathogen (Phlyctema padi or formerly Cylindrosporium padi) has an asexual fruiting body called an acervulus that forms a gelatinous white tendril (cirrhus) of conidia 

within the center of the leaf lesion during moist weather (fig. 4-4b). Conidia are splash 

dispersed by rain and wind and cause secondary infections throughout the summer. Fresh 

pruning wounds can also become infected. Warm temperatures are especially conducive 

to disease development. Heavy infection may cause premature leaf fall; some trees may 

be completely defoliated by July. 

Shot-hole blight—Infection by S. carpophila begins in spring and is favored by cool, 

wet springs. The pathogen overwinters in blasted buds and cankers, and the primary 

inoculum generally originates from such tissues, as sporulation occurs during wet spring 

weather. Fall applied protectant fungicides protect the buds from fungal infection during 

the late and dormant seasons. Peach (Prunus persica) and apricot (Prunus armeniaca) are 

more susceptible to S. carpophila than cherry, but the pathogen can infect most species of 

Prunus.

Cercospora leaf spot—Leaf spots and shot-hole symptoms caused by M. cerasella are 

favored by wet springs. The pathogen overwinters on fallen leaves and produces multiple 

sexual fruiting bodies in each former leaf spot. The primary inoculum is the ascospores, 

which are discharged in air currents during spring and infect leaves with wet surfaces 

during moist weather. The summer form of the pathogen (Passalora circumscissa, formerly Cercospora circumscissa) is a stroma with fascicles of conidiophores and conidia. 

Conidia that form in the necrotic leaf lesions become windblown and cause secondary 

infections throughout the summer during conducive weather conditions. The pathogen is 

common on other species of Prunus.

Damage

Cherry leaf spotting diseases cause defoliation; severe disease reduces production and 

storage of carbohydrates and causes reduced growth. Repeated infections and early defoliation alter the condition of the trees going into winter dormancy and reduce the number 

of leaves formed and the fruit yield the next year. The reduced vigor the following spring 

sometimes can result in mortality. Additionally, diseased trees become more sensitive to 

bud mortality after frost injury. Trees with repeated infections decline over several years.

Cherry leaf spot caused by B. jaapii greatly damages commercial sweet and sour 

cherries in parts of the United States. Although effects on Nanking cherry, chokecherry,

and sandcherry are less dramatic, the pathogen commonly causes severe defoliation in 

crowded nursery beds of chokecherry and sandcherry under sprinkler irrigation in Great 

Plains nurseries. Planting of chokecherry has been reduced in parts of the Great Plains 

because of this disease. 

Fungal shot-hole blight from repeated infections by S. carpophila causes considerable branch dieback owing to the accumulation of bud mortality (blasting) and annual 

twig cankers. As a result, fruit-bearing and flowering branches are lost, and aesthetic 

value and vigor are diminished.

Bacterial leaf spot and shot-hole blight caused by X. arboricola pv. pruni is less 

severe on cherry than other stone fruits, and annual cankers and shoot and twig dieback 

are not usually present on cherry. The perennial deep-seated cankers that occur on plum 

(Prunus spp.) do not occur on cherry. This disease is the most economically damaging 

because of the severe infection of fruit. When fruit production is not the objective of 

planting, the disease becomes of little significance on cherry except in the nursery, where 

management of shot-hole is problematic. 
USDA Forest Service RMRS-GTR-335. 2016. 25

P. syringae pv. syringae is more important as a canker-causing pathogen of residential 

and commercial stone fruit trees than as a leaf spot pathogen. Large perennial cankers 

on the trunk and major scaffold branches produce copious gummosis and are highly destructive. The effect of this pathogen on Nanking cherry, sandcherry, and chokecherry is 

limited, although some windbreak and ornamental plantings may be severely infected. 

Management

Usually, managing these leaf spot diseases is necessary only in commercial fruit growing areas and in nurseries. B. jaapii can be controlled with spring curative fungicide and 

summer protectant fungicide treatments applied directly to foliage. An epidemiological 

model of the disease cycle is available, which aids in predicting periods of high disease 

probability in commercial cherry orchards (Eisensmith and Jones 1981). The number 

of fungicide applications necessary to control the disease has been reduced by use of 

curative fungicides for spring application to decrease ascospore infection, and use of this 

predictive model (Holb and others 2010).

Leaf spot caused by S. carpophila, X. arboicola pv. pruni, and P. syringae pv. syringae

can be reduced by incorporating sanitation practices, such as dormant pruning of budblasted and cankered wood and removal of leaves and other debris in the fall. Dormant 

pruning of cankered branches is especially important for P. syringae pv. syringae. 

Fungicides can be used to suppress disease development of S. carpophila in the season 

and also during dormancy. Bactericides and fixed copper applications can decrease the 

damage of the bacterial diseases, but use copper compounds only in the early season as 

they can blemish fruit destined for market. Best disease control is obtained with several 

foliar applications of fungicides or bactericides coordinated with periods of high infection 

hazard. Fungicides or bactericides alone do not provide good control. A combination of 

sanitation and fungicide application is recommended.

Selected References

Battilani, P.; Rossi, V.; Saccardi, A. 1999. Development of Xanthomonas arboricola pv. 

pruni epidemics on peach. Journal of Plant Pathology. 81: 161–171.

Cameron, H.R. 1962. Mode of infection of sweet cherry by Pseudomonas syringae. 

Phytopathology. 52: 917–921.

Eisensmith, S.P.; Jones, A.L. 1981. A model for detecting infection periods of 

Coccomyces hiemalis on sour cherry. Phytopathology. 71: 728–732.

Holb, I.J.; Lakatos, P.; Abonyi, F. 2010. Some aspects of disease management of cherry 

leaf spot (Blumeriella jaapii) with special reference to fungicide use. International 

Journal of Horticultural Science. 16(1): 45–49. 

Jenkins, W.A. 1930. The cherry leaf-spot fungus, Mycosphaerella cerasella Aderh. Its 

morphology and life history. Phytopathology. 20: 329–337.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Zehr, E.I.; Shepard, D.P. and Bridges Jr., W.C. 1996. Bacterial spot of peach as influenced 

by water congestion, leaf wetness duration, and temperature. Plant Disease. 80: 

339–341.
26 USDA Forest Service RMRS-GTR-335. 2016.

5. Chlorosis 

Laurie J. Stepanek, Mark O. Harrell, and Rachel A. Allison; revised 

from Mark O. Harrell and Mark W. Andrews (Riffle and Peterson 1986)

Chlorosis describes any condition in which leaves or needles develop an abnormally 

light green or yellow color. The most common cause of chlorosis in trees is a deficiency 

of usable iron, either in the soil or within the tree. But the problem is complex, often 

involving multiple factors, and can be difficult to correct.

Hosts and Distribution

Chlorosis may develop wherever trees are grown in alkaline soils (soils with a pH 

greater than 7). Alkalinity generally increases from east to west across the Great Plains, 

and with it, incidence of chlorosis. Chlorosis is also common in urban areas, where excavation during construction of buildings brings alkaline subsoil to the surface and where 

poor cultural practices lead to unhealthy soils and root damage in trees.

Pin oak (Quercus palustris) and silver maple (Acer saccharinum) are commonly affected by chlorosis, but many other tree species are susceptible as well (table 5-1). Trees 

susceptible to chlorosis differ somewhat among the various areas of the Great Plains.

Table 5-1—Partial list of trees susceptible to chlorosis.

 Common name Scientific name Common name Scientific name

Amur maple Acer ginnala Crabapple Malus spp.

Red maple A. rubrum Pine Pinus spp.

Silver maple A. saccharinum Peach Prunus persica

Freeman maple A. x freemanii Pear Pyrus spp.

Serviceberry Amelanchier spp. Aspen Populus spp.

River birch Betula nigra Cottonwood Populus spp.

Catalpa Catalpa speciosa Poplar Populus spp.

Flowering dogwood Cornus florida Swamp white oak Quercus bicolor

Hawthorn Crataegus spp. Pin oak Q. palustris

Holly Ilex spp. Northern red oak Q. rubra

Juniper Juniperus spp. Baldcypress Taxodium spp.

Sweetgum Liquidambar styraciflua Yew Taxus spp.

Magnolia Magnolia spp.

Symptoms and Signs

The degree of yellowing varies from a yellowish-green in only slightly chlorotic 

leaves to lemon yellow and almost white in leaves that are moderately to severely chlorotic (figs. 5-1 and 5-2). The yellowing is most intense in the interveinal areas of leaves 

while the veins remain green. Brown necrotic spots may develop between the veins and 

along the leaf edges, giving the appearance of leaf scorch (fig. 5-3), and leaves may be 

abnormally small.

Symptoms of chlorosis may be uniform throughout the entire tree or be confined to 

one or a few branches. If chlorosis continues for several years, shoot growth is reduced, 

branches begin to die back, and the tree may eventually die (fig. 5-4). Symptoms among 

trees of the same species growing on the same site can vary; even normal green trees 

can be located next to severely chlorotic ones owing to soil variations within the site and 

genetic differences among trees (fig. 5-5).
USDA Forest Service RMRS-GTR-335. 2016. 27

Figure 5-3—Necrotic spots, which may develop 

along leaf edges and between the veins (Laurie 

J. Stepanek, Nebraska Forest Service, used 

with permission).

Figure 5-1—Chlorotic Freeman maple leaves 

showing typical yellow interveinal areas and 

green veins (Laurie J. Stepanek, Nebraska 

Forest Service, used with permission).

Figure 5-2—Chlorotic baldcypress foliage (Laurie 

J. Stepanek, Nebraska Forest Service, used with 

permission).

Figure 5-4—Branch dieback in a chlorotic crabapple. Symptoms developed shortly after an 

irrigation system was installed and the landscape 

was overwatered (Rachel A. Alison, Nebraska 

Forest Service, used with permission).

Figure 5-5—Variation in symptoms among oak 

trees (Mark O. Harrell, Nebraska Forest Service, 

used with permission).

Cause

Deficiency of iron is the most common 

cause of chlorosis of trees in the Great 

Plains. In most cases, iron in the soil is 

present in sufficient quantity, but under 

alkaline conditions, it becomes less soluble 

and thus less available for plant uptake. 

Even if absorbed, iron may remain in a 

form that the tree is unable to use. Iron is 

used in the production of chlorophyll, so 

a deficiency of iron prevents the leaves 

from producing the normal amount of 

chlorophyll.

The availability of other micronutrients such as manganese and zinc also is limited in 

alkaline soils and therefore may lead to chlorosis as well. An excess of some nutrients, 

especially nitrate-nitrogen and phosphorus, can cause chlorosis by interfering with 

the uptake of iron in the soil or its movement and utilization in the tree. High levels of 

bicarbonates, such as those found in calcareous soils and many sources of irrigation 

water and groundwater, interfere with these processes as well. A lack of nitrogen or other 

macronutrients may cause chlorosis, but the focus of this discussion is on micronutrient 

deficiencies associated with high pH soils.

Chlorosis frequently develops in wet, compacted soils, such as those found in overwatered landscapes (fig. 5-4). Saturated soils limit gas exchange, allowing the buildup of 

carbon dioxide, which leads to increased bicarbonate levels. Extensive rains in spring can 
28 USDA Forest Service RMRS-GTR-335. 2016.

cause temporary chlorosis. Planting trees too deeply, piling soil over roots, and digging or 

trenching near trees can damage roots, resulting in limited nutrient uptake and the development of chlorotic tissues.

Damage

Trees with severe chlorosis typically decline and die over several years. In many areas 

of the Great Plains some species, especially pin oak, are no longer being planted because 

this disorder severely reduces their chances of survival.

Management

Chlorosis is difficult to correct. Soils resist changes in pH, and several factors that 

contribute to the chlorotic condition may be involved. Treatments that work in one area 

may not work in other areas, and some trees respond better to certain treatments than 

others. A general discussion of control options is provided here. Consult your local university extension office, state forestry department, or the U.S. Forest Service for specific 

information.

Cultural practices should be included in any treatment plan. Water trees thoroughly but 

infrequently if it does not rain, which will give time for the soil to drain and allow gas exchange to occur. Automatic sprinklers that run daily or every other day can severely stress 

or kill trees. Woodchip mulches improve the soil environment for tree roots, mycorrhizal 

fungi, and other beneficial soil microbes that aid in nutrient uptake, as well as provide 

a natural supply of micronutrients. Use fertilizers with prudence. Trees generally do not 

need extra nitrogen fertilizer if the lawn is already being fertilized.

When planting trees, choose those that are more adapted to alkaline soils. Specific 

trees will vary widely across the Great Plains so consult local sources for recommendations. Avoid pot-bound trees with circling roots and prepare a wide, shallow planting site 

(not a deep hole) to keep the main roots level with the soil surface. Remove all twine, 

wire, and burlap and spread the roots out as much as possible, which will help the tree 

establish a healthy root system.

Several chemical treatments are available to help correct chlorosis, including soil 

applications, trunk injections or implants, and foliar sprays. Soil and leaf tissue tests can 

provide information on pH and nutrient levels, which can help with treatment selection.

Soil treatments

Micronutrient deficiency in soils is uncommon. The goal of most soil treatments, 

therefore, is to lower the soil pH to improve nutrient availability. Elemental sulfur is commonly used to achieve this. Soil bacteria convert the sulfur to acid, which lowers the pH. 

Results are variable and slow to work (symptoms may not improve for one to two years 

after treatments), but the effects can last several years.

A broadcast application of sulfur can be applied below the canopy of the tree. High 

rates are needed: one to five pounds or more per 100 ft2, depending on initial pH and soil 

texture. Raking the sulfur into mulched areas and aerating turf areas will increase sulfur/

soil contact and speed up the acidification process. Watering thoroughly after application 

and during hot, dry weather will help prevent turfgrass burn. Annual followup applications of sulfur can help maintain desired pH levels. Broadcast applications may be less 

successful on clay soils, which are highly buffered and more resistant to pH changes than 

sandy soils.

Alternatively, sulfur may be combined with iron sulfate or a micronutrient mix and 

placed into holes dug in the ground around the tree. Pockets of soil are created that are 

rich in readily available iron and other micronutrients. This method may be a better option for clay soils.

Calcareous soils contain high levels of free lime (calcium carbonate) and are extremely difficult to correct. Excessive amounts of sulfur are needed to neutralize the free 

lime before the pH can be lowered. To test for free lime, moisten a small amount of dry 

soil with vinegar. If the mixture fizzes or bubbles, it has free lime.

Another soil-applied chemical specifically for trees deficient in iron is iron chelate, 

which provides iron in a form easily absorbed by plants. Look for products containing 
USDA Forest Service RMRS-GTR-335. 2016. 29

EDDHA (FeEDDHA), which works better than other chelates when soil pH is above 7.2. 

This chelate has been shown to be effective even in calcareous soils. Iron chelates generally provide control for one to two years.

Trunk injections and implants

In areas where soil treatments may be ineffective or impractical, trunk injections and 

implants are commonly used to treat chlorosis. For injections, holes are drilled into the 

trunk flare, and a liquid micronutrient product is injected into the holes; or the product 

is delivered to the tree through a needlelike injection tool. Implants are plastic cartridges 

containing a powdered material and are placed into holes drilled into the trunk. Most 

injections and implants are effective for one to three years.

One major disadvantage of injections and implants is the physical damage caused by 

the drilling and the internal damage caused by the chemical itself. Treatments should not 

be re-applied until the drill wounds seal over with new trunk tissue, which may take more 

than a year. Injection methods that use small, shallow holes are less damaging. Although 

some products are available to consumers, the potential for failure and significant tree 

injury is high. Injection treatments performed by a certified arborist who is well trained in 

the procedure can reduce these risks.

Spray treatments

Spray treatments for chlorosis consist of a solution of iron or manganese sulfate or a 

chelated compound. This treatment improves the condition of only those leaves present 

when the foliage is sprayed. It has little or no effect on leaves formed after spraying, and 

the effectiveness does not carry over into the next growing season. Used in combination 

with a soil treatment, a spray can provide iron to the tree until the slower acting soil treatment becomes effective.

Selected References

Barton, L.L.; Abadía, J., eds. 2006. Iron nutrition in plants and rhizospheric 

microorganisms. Dordrecht, the Netherlands: Springer. 477 p.

Harrell, M.O.; Pierce, P.A.; Mooter, D.P. 1988. Pin oak and silver maple chlorosis 

treatment with ferric ammonium citrate solution. Journal of Arboriculture. 14(6): 

156–158.

Harrell, M.O.; Pierce, P.A.; Mooter, D.P.; Webster, B.L. 1984. A comparison of treatments 

for chlorosis of pin oak and silver maple. Journal of Arboriculture. 10(9): 246–249.

Korcak, R.F. 1987. Iron deficiency chlorosis. Horticultural Reviews. 9: 133–186.

Mengel, K. 1995. Iron availability in plant tissues–iron chlorosis on calcareous soils. In: 

Abadía, J., ed. Iron nutrition in soils and plants. Dordrecht, the Netherlands: Kluwer 

Academic Publishers: 389–397. 

Neely, D. 1976. Iron deficiency chlorosis of shade trees. Journal of Arboriculture. 2: 

128–130.

Neely, D. 1980. Trunk and soil chlorosis treatments of pin oak. Journal of Arboriculture. 

6(11): 298–299.

Whitcomb, C.E. 2006. Establishment and maintenance of landscape plants II. Stillwater, 

OK: Lacebark Inc. 340 p.
30 USDA Forest Service RMRS-GTR-335. 2016.

6. Herbicide Damage to Trees and Shrubs

Aaron D. Bergdahl and William R. Jacobi

Intensified use of herbicides in recent decades has increased the hazard to nontarget 

vegetation. Disorders or mortality caused by herbicides is particularly prevalent among 

sensitive tree species in residential and agricultural settings where herbicides are 

commonly used. Areas at higher risk for exposure include windbreaks, sites near rightsof-way, railroads, roadsides, or areas treated for noxious weed control.

Herbicide Classification and Modes of Action

Herbicides that commonly cause injury to nontarget plants fall into three general 

groups: 1) those that are used to kill a certain type of established vegetation (post-emergence selective herbicides), 2) herbicides intended to prevent germination of seeds or 

emergence of seedlings (preemergence selective herbicides) or 3) those that kill all vegetation (nonselective herbicides). Herbicides are classified as either contact herbicides, 

which affect only the points of contact with the active ingredient, or systemic herbicides, 

which are absorbed, are translocated, and typically show a broader pattern of symptoms. 

Herbicides kill plants by interfering with essential plant functions; this interference 

causes disruption of cell components and inhibits proteins essential to photosynthesis, 

growth, and cell formation. The modes of action of herbicides most commonly found to 

damage trees in the Great Plains are described here.

Plant growth regulator herbicides are typically used on turf to selectively kill broadleaf 

weeds. Herbicides in this group include the phenoxyacetic acids (2,4-D, 2,4-DP, MCPA, 

MCPP, and related compounds), benzoic acids (for example, dicamba), and pyridine carboxylic acids (for example, clopyralid, fluroxypyr, picloram, and triclopyr). Herbicides 

of this type can move systemically to growing points, where they disrupt plant growth 

Figure 6-1—(A) Phenoxy-type herbicide injury to green ash (Aaron D. Bergdahl, North Dakota 

Forest Service, used with permission); (B) Simazine damage on poplar (John Sharpe, Prairie 

Farm Rehabilitation Administration, Indian Head, Saskatchewan, used with permission); (C) 2,4-D 

damage on maple (Robert Stack, North Dakota State University, retired, used with permission); 

(D) Glyphosate plus imazapyr damage on linden (Michael Schomaker, Colorado Forest Service, 

retired, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 31

hormone balance, causing various developmental deformities such as narrow elongated 

growth (leaf strapping), cupping, and twisting (figs. 6-1a, c, 6-2, 6-3, 6-5, and 6-7a and b). 

Exposure to thin barked species by these chemicals may result in deformation, blistering, and cankering of bark (fig. 6-4). Symptoms develop within a few days to a week 

after exposure. Pyridine carboxylic acid herbicides can remain active in the environment 

(residual period) for more than one year. 

Figure 6-2—Symptoms of 2,4-D damage showing twisted new growth and cupped leaves on (A) linden, and (B) elm (Aaron D. Bergdahl, North Dakota Forest Service, used with permission).

Figure 6-3—Strapping and discoloration of bur 

oak (Quercus macrocarpa) foliage resulting from 

herbicide exposure (Aaron D. Bergdahl, North 

Dakota Forest Service, used with permission).

Figure 6-4—Blistering of bark on honeylocust from herbicide exposure (William R. 

Jacobi, Colorado State University, used with 

permission).

A B
32 USDA Forest Service RMRS-GTR-335. 2016.

Figure 6-5—Picloram damage to catalpa (Catalpa speciosa) (Ned Tisserat, Colorado State University, used with 

permission).

Figure 6-6—Glyphosate damage to red oak (Quercus 

rubra) and adjacent white spruce (Picea glauca) (Aaron 

D. Bergdahl, North Dakota Forest Service, used with 

permission).

Commonly used nonselective herbicides belonging to the group known as photosynthesis inhibitors include the following (with examples in parentheses): triazines (atrazine, 

simazine), phenylureas (tebuthiuron), uracils (terbacil), benzothiadiazoles (bentazon),

nitriles (bromoxynil), and carbamates (desmedipham). These herbicides cause bleaching 

of leaves followed by leaf browning and drying out (fig. 6-1b). Effects may be exaggerated in higher-pH soils and the residual period may be several months. 

Nonselective herbicides, also called bare ground or total vegetation control herbicides, 

function via a variety of modes of action, ranging from interfering with amino acid 

formation to disrupting lipid formation (plant cell membrane disruption). Nonselective 

herbicides like glyphosate and bromacil are applied directly onto actively growing 

target plants. The development of crop varieties that are glyphosate resistant has led to 

greatly expanded use of these compounds in recent years, resulting in increased reports 

of glyphosate damage to trees, especially in field windbreaks (figs. 6-1d and 6-6). 

Nonselective herbicides may have long-lasting residual activity, so effects can be persistent. Some products also include multiple chemicals and therefore multiple modes of 

action, creating variable symptoms that are not as diagnostic. 

Preemergence herbicides are applied directly to soil or on turf and then incorporated 

into the soil, usually with water, to kill germinating weed seedlings. Preemergence 

herbicides are less commonly associated with tree disorders in the northern Great Plains, 

mainly because of the mode and timing of application.

Symptoms

Symptoms vary depending on the type and concentration of the chemical, frequency 

of use, species and condition of plants, and the timing of chemical application. The 

more common expressions of injury from phenoxy or hormonal-type herbicides include 

cupped leaves, parallel leaf venation on normally net-veined leaves, 

chlorosis, nastic growth, and wavy 

or curled leaf margins (figs. 6-1a 

and 6-1c, 6-2, and 6-5). Lindens 

(Tilia spp.), honeylocust (Gleditsia 

triacanthos), redbud (Cercis 

canadensis), and boxelder (Acer 

negundo) are among those that 

readily exhibit these effects.

Leaves of some trees, such 

as pin oak (Quercus palustris), 

become waxy and stiff, while 

maples may develop pebbled foliage that appears weather-beaten. 

Loss of apical growth is typical 

of phenoxy or hormonal herbicide 

injury. Affected trees may suffer a 

gradual crown dieback and eventually die. Lateral leaf development 

may also be hindered. Exposure 

to 2,4-D may result in the production of fewer than normal leaves, 

flowers, and fruits. Leaf scorch can 

indicate herbicide injury, particularly when exposure is heavy. Ash 

(Fraxinus spp.) and cottonwood 
USDA Forest Service RMRS-GTR-335. 2016. 33

Figure 6-7—Examples of herbicide damage to evergreens. (A) Spiral pattern typical of a point exposure to herbicide in spruce; Aaron D. Bergdahl, North Dakota Forest Service, used with permission), 

(B) MCPA direct exposure to Scots pine (Pinus sylvestris) (James A. Walla, Northern Tree Specialties, 

used with permission); (C) Spruce injury from soil sterilant exposure (William R. Jacobi, Colorado 

State University, used with permission).

(Populus deltoides) may exhibit scorched leaves without expressing broader effects of 

herbicide exposure. High rates of exposure of 2,4-D to Siberian elm (Ulmus pumila) have 

been shown to result in bark abnormalities. A single 2,4-D exposure may produce injury 

for two years or more. Honeylocust bark can become spongy, woody swellings (galls) 

can form at branch crotches, and the cambium may be killed by high concentrations of 

phenoxy herbicides (fig. 6-4).

Evergreens are generally resistant to phenoxy herbicides, but they may be injured 

under certain circumstances. On conifers symptoms can include abnormal swelling, twisting, and curling of branchlets, as well as bleaching, browning, and casting of needles. 

Broadleaf trees and especially conifers have shown sensitivity to dicamba often used in 

“weed and feed” lawn applications. Dicamba can be absorbed via the roots and injury to 

trees ranges from leaf distortion, discoloration, and defoliation to branch dieback, often 

expressed in a spiral pattern in conifers (fig. 6-7a), and sometimes tree mortality. When 

farm-grade broadleaf weed control products that frequently combine more than one 

herbicide mode of action are applied to lawns, they may cause tree mortality in landscape 

settings, especially on lighter soil types.

Nonselective herbicides, which are used for bare ground or total vegetation control, 

tend to halt growth and cause chlorosis of new and old foliage. At high doses, foliar 

browning, foliar drop, and dieback of twigs and branches may occur (fig. 6-7c). Trees 

and shrubs injured by these herbicides are less likely to recover than trees injured by 

plant growth regulator herbicides. While light exposure to some herbicides may not cause 

noticeable symptoms or damage, partial or total mortality can result from exposure to 

nonselective herbicides.

Diagnosis of herbicide damage can be aided by careful collection and analysis of 

residue samples. The cost of analysis is usually prohibitive, but if analysis is required, 

the testing laboratory will need to know what herbicides to test for in the plant samples. 

It is important that the samples be collected as soon as possible after exposure. When 

possible, residue samples should be kept frozen until they reach the laboratory. It may be 

advisable to contact an analytic laboratory for specific instructions on collection and care 

of samples.
34 USDA Forest Service RMRS-GTR-335. 2016.

Damage

All possible sources of chemical exposure should be considered when dealing with 

suspected herbicide damage. Harmful exposure to herbicides can result from drift of the 

spray particles, movement of volatiles, application to the soil or movement in soil (or 

water) resulting in root exposure, and direct application. Spray drift volatiles can move 

considerable distances, resulting in damage several miles from where the herbicide was 

applied. Direct application may cause damage if sensitive trees are not carefully avoided.

In urban areas, desirable plants are most likely to be damaged by repeated lawn sprays 

or fertilizers that contain herbicides. Damage is also commonly associated with soil

applied herbicides used around driveways, sidewalks, and structures to remove vegetation. In situations where damage has resulted from an apparent misuse of herbicides, it 

may be appropriate to contact your state’s pesticide regulatory agency or department of 

agriculture for guidance.

Management

Desirable trees located where herbicide exposure is likely will require protection. 

Persons who apply herbicides should be informed of locations of desirable trees, and 

products that pose a minimal risk to the trees should be used. Exercise extra caution to 

ensure proper calibration of equipment and application. Herbicides that are mishandled or 

are applied under improper conditions often end up injuring nontarget plants. Avoid misapplication to nontarget plants by applying sprays only when it is not windy so that drift 

of herbicide will be unlikely. Additionally, vapor drift can occur with many plant growth 

regulator herbicides if temperatures are too high during application or soon afterwards.

When applying broadleaf herbicides to turf, remember that tree roots can extend up to 

five times beyond the diameter of the tree crown and herbicides can be absorbed by tree 

roots present in the area of application. 

Once exposure has occurred to plants, little can be done to minimize the initial effects. 

Washing herbicide off foliage can be effective if it is done immediately after exposure. 

Soil-active herbicides can be deactivated on a limited scale with activated charcoal or 

similar products. Generally, any effort that promotes tree vigor should help reduce the 

effects of the herbicide exposure. Trees weakened by herbicides may be predisposed to 

insects, pathogens, and other types of environmental damage. Many trees can survive 

moderate herbicide damage if given care after exposure. Trees should be given at least 

one year to recover before declaring the tree not salvageable. Inappropriate and high, 

acute dosages of herbicides can kill trees, especially if the trees are stressed from growing 

under droughty conditions along roadsides or in windbreaks. Always check with your 

state’s pesticide regulatory agency for current registered use of pesticides and strictly follow all aspects of product labels.

Selected References

Gunsolus, J.L.; Curran, W.S. 1999. Herbicide mode of action and injury symptoms. North 

Regional Publ. 377. St. Paul, MN: University of Minnesota Extension. 24 p.

Hibbs, R.H. 1978. Recognition of weed-killer injury to trees. Journal of Arboriculture. 4: 

189–191. 

Otta, J.D. 1974. Effects of 2,4-D herbicide on Siberian elm. Forest Science. 20: 287–290.

Sherwood, C.H.; Weigie, J.L.; Denisen, E.L. 1970. 2, 4-D as an air pollutant; effects on 

growth of representative horticultural plants. Horticultural Science. 5:202, 211–213. 

Tu, M.; Hurd, C.; Randall, J.M. 2001. Weed control methods handbook: tools and 

techniques for use in natural areas. April 2001 version. The Nature Conservancy. 

http://www.invasive.org/gist/handbook.html. (February 25, 2015).
USDA Forest Service RMRS-GTR-335. 2016. 35

7. Taphrina Diseases of Shade and Fruit Trees

Mark L. Gleason, Hayley M. Nelson, and Megan M. Kennelly

Species of fungi in the genus Taphrina infect a wide range of woody plants, but primarily oak (Quercus spp.), maple (Acer spp.), elm (Ulmus spp.), plum (Prunus spp.), and 

peach (Prunus persica) in the Great Plains. These fungi cause conspicuous blistering and 

browning of leaves as well as distortion of fruit and defoliation. 

Hosts and Distribution

Species of fungi in the genus Taphrina are host specific, meaning that each one infects 

only certain tree species. Taphrina caerulescens infects many oaks, primarily those in 

the red oak group: red (Q. rubra), scarlet (Q. coccinea), black (Q. velutina), and pin oak 

(Q. palustris). Sometimes it also infects those in the white oak group: bur (Q. macrocarpa) 

and white (Q. alba) oak. Within T. caerulescens there are subgroups that are specific to 

certain types of oaks.

Taphrina carveri (named after the famous plant scientist George Washington Carver, 

who originally described its sexual spores) attacks maple species, primarily red (A. 

rubrum) and silver (A. saccharinum) maple, and T. ulmi attacks elm leaves. Both T. 

communis and T. pruni cause plum pockets on plum fruit, but T. communis is much more 

prevalent in the Great Plains. A number of cultivated and wild plums are susceptible, but 

cultivated plums are less so. Peach leaf curl, caused by T. deformans, can be a damaging 

disease in commercial and backyard peach and nectarine (Prunus spp.) orchards. 

Symptoms and Signs

Most Taphrina diseases attack 

leaves, and symptoms typically appear 

in late spring to early summer. Oak 

leaf blister appears on the upper leaf 

surfaces as circular, raised, wrinkled, 

yellowish-white spots, and on the leaf 

undersides as yellowish-brown to gray 

depressions of the same size directly 

below the raised areas (figs. 7-1 and 

7-2). The blistered tissue later turns 

reddish brown with yellow margins, 

and finally becomes dull brown. When 

several blisters merge, leaves may 

become curled and fall off prematurely. 

Maple leaf blister produces grayishbrown to black, irregularly shaped 

spots that are sometimes slightly raised. 

As the leaf spots age, their centers 

often turn a lighter color (fig. 7-3). 

Multiple infections cause the leaf shape 

to become distorted (fig. 7-4). The leaf 

spots’ somewhat rounded shape helps 

to distinguish them from spots caused 

by maple anthracnose, which tends to 

produce irregularly shaped, angular 

leaf spots. Elm leaf blister causes yellow spots that enlarge into dull brown, 

somewhat angular spots with light 

green halos (fig. 7-5) and some areas of 

puckered, light green tissue (fig. 7-6).

Figure 7-1—Leaf blistering and discoloration caused 

by oak leaf blister (Cornell University, used with 

permission).

Figure 7-2—Yellow and brown discoloration caused 

by oak leaf blister (Cornell University, used with 

permission).
36 USDA Forest Service RMRS-GTR-335. 2016.

Figure 7-3—Coalescing leaf spots caused by maple 

leaf blister (Sabrina Tirpak, Plant Diagnostic Lab, 

Rutgers University, used with permission).

Figure 7-4—Figure 4. Distortion caused 

by maple leaf blister and anthracnose. 

(Gail Ruhl, Purdue University, used with 

permission).

Figure 7-5—Leaf spots caused by elm leaf blister (Megan Kennelly, Kansas State University, 

used with permission).

Figure 7-6—Blistering and browning of leaf 

caused by elm leaf blister (Megan Kennelly, 

Kansas State University, used with permission).

Symptoms of plum pockets are most noticeable on fruit. Small, whitish spots appear 

on young fruit, then expand in size to cover the fruit. Seed fails to form in the infected 

fruit, and the fruit become hollow and enlarge to many times their normal size (fig. 7-7). 

They sometimes take on a red to grayish cast and a thickened, leathery texture (fig. 7-8). 

Peach leaf curl results in puckered to severely distorted, downward-curling leaves 

that are crisp in texture and show red and purple discoloration (fig. 7-9). Later, the leaf 

surface turns grayish as the fungus begins to produce spores. Diseased leaves eventually 

die, fall off the tree, and are replaced by a new crop of leaves in early to midsummer. 

Extensive defoliation can lead to a reduced fruit crop the following year. Though less 

apparent, fruits and flowers can also be directly infected, often falling off the tree. Young 

shoots can also be stunted and deformed.

Disease Cycle

Taphrina fungi overwinter as dormant spores in bud scales and bark crevices. These 

spores germinate in spring during cool, rainy periods and infect young leaves and fruit. 

The spores are produced on infected tissues and later spread by wind and rain, but do not 

cause additional infections during the same growing season. Instead, surviving spores 

remain dormant until favorable weather occurs the next spring. As a result, Taphrina

diseases have only one cycle of infection per year.

Damage

Damage to trees is primarily cosmetic, and most Taphrina diseases do not endanger 

plant health. On peach and nectarine, however, defoliation by peach leaf curl can weaken 

trees and reduce yield if not controlled.
USDA Forest Service RMRS-GTR-335. 2016. 37

Management

Most Taphrina diseases do not need to be controlled because their threat to tree health 

is negligible. The exception is in peach and nectarine orchards, where peach leaf curl can 

cause economic losses; in those situations, a single fungicide spray, applied between the 

time that 90 percent of leaves have been shed in fall and the time that leaf buds begin to 

swell in spring, is highly effective at controlling the disease. Appropriate water and fertilizer may reduce stress and promote tree vigor in affected trees. Although full resistance is 

not available, some peach cultivars have reduced susceptibility. 

Selected References

Jensen, S.L. 2013. Oak leaf blister (Taphrina caerulescens). Ithaca, NY: Cornell 

University. http://plantclinic.cornell.edu/factsheets/oakleafblister.pdf. (February 25, 

2015).

Hartman, J.; Bachi, P. 2004. Peach leaf curl and plum pockets. Bull. PPFS-FR-T-01, 

Lexington, KY: University of Kentucky. 2 p. http://www2.ca.uky.edu/agcollege/

plantpathology/ext_files/PPFShtml/PPFS-FR-T-1.pdf. (February 25, 2015).

Figure 7-7—Plum fruit infected by plum 

pockets is yellow and abnormally enlarged 

(Thomas Whitlow, Cornell University, used with 

permission).

Figure 7-8—Gray discoloration of plum fruit 

caused by spores of the plum pockets fungus 

(Aaron Bergdahl, North Dakota Forest Service, 

used with permission).

Figure 7-9—Discoloration and puckering caused by peach leaf curl (Sabrina Tirpak, Plant 

Diagnostic Lab, Rutgers University).
38 USDA Forest Service RMRS-GTR-335. 2016.

8. Leaf Tatter and Leaf Tatters 

Laurie J. Stepanek and Mark O. Harrell

Leaves of broadleaf trees sometimes sustain various types of abiotic injury that cause 

the leaves to appear lacy or tattered. This tattering condition has been referred to by several similar terms, which are sometimes used interchangeably. “Tatter” (also “leaf tatter” 

and “tatter-leaf”) appears in early scientific literature and service publications, where it 

is broadly applied to tattering symptoms that are associated with unknown abiotic causes 

and that affect a variety of tree species. The similar term “tatters” (or “leaf tatters”) is 

found in more recent research and describes damage caused by specific herbicides on a 

limited number of species.

In this discussion, “leaf tatter” will refer to a general abiotic tattering disorder without regard to cause or host, and “leaf tatters” will be used exclusively for the specific 

herbicide-induced disorder affecting oak (Quercus spp.) and common hackberry (Celtis 

occidentalis).

Hosts and Distribution

Leaf tatter occurs on many broadleaf trees throughout the Great Plains, including 

species of maple (Acer), oak, linden (Tilia), birch (Betula), ash (Fraxinus), crabapple 

(Malus), hawthorn (Crataegus), dogwood (Cornus), and hackberry. Maple species, 

especially sugar maple (A. saccharum), are particularly prone to leaf tatter. Susceptible 

cultivars include ‘Green Mountain®,’ ‘Wright Brothers,’ ‘Bonfire,’ ‘Endowment,’ and 

‘Fairview.’

Leaf tatters, the herbicide-induced condition of tattering, occurs on hackberry and 

oak, particularly bur (Q. macrocarpa), white (Q. alba), and northern red (Q. rubra) oak. 

Leaf tatters occurs principally in midwestern states but is known to occur in the eastern 

Great Plains. Many cases occur in rural areas or near crop fields where agricultural herbicides are used. Native stands of trees as well as planted ornamental trees can be affected.

Symptoms

Leaf tatter may show a range of symptoms. One symptom found in many species is 

the presence of numerous holes between the veins (fig. 8-1). The holes may extend to the 

margins, affecting the form of the leaf and creating a ragged appearance.

Other symptoms, often seen in maples, are shredding, tearing, or absence of tissue between the lobes of the leaves (fig. 8-2). The tips of the lobes may develop a water-soaked 

appearance, followed by browning, drying, and disintegration of the tissues. The palmate 

shape typical of maple leaves may be lost when tatter is extensive (fig. 8-3).

Figure 8-1—Leaf tatter on birch associated with a late spring freeze (Laurie 

J. Stepanek, Nebraska Forest Service, 

used with permission).

Figure 8-2—Leaf tatter of sugar maple 

showing missing tissue between the 

lobes (Laurie J. Stepanek, Nebraska 

Forest Service, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 39

Figure 8-3—Sugar maple with missing leaf tips resulting in loss of palmate 

shape (Laurie J. Stepanek, Nebraska 

Forest Service, used with permission).

Figure 8-4—Bur oak leaf with tatters. Dead tissue 

falls away, leaving mostly veins (Laurie J. Stepanek, 

Nebraska Forest Service, used with permission).

Figure 8-5—Hackberry leaves with 

varying degrees of damage from tatters 

(Laurie J. Stepanek, Nebraska Forest 

Service, used with permission)

Figure 8-6—Bur oak leaves reduced to veins 

(Laurie J. Stepanek, Nebraska Forest Service, 

used with permission).

Symptoms of leaf tatters on oak and hackberry appear on young leaves as they unfold 

and begin to expand in spring. Leaf edges and interveinal areas turn brown, and leaves 

may curl or pucker. As the damaged parts of the leaves die and drop off, the leaves develop a lacy appearance (fig. 8-4). Leaf damage can vary from only a few scattered holes to 

leaves that are little more than veins (figs. 8-5 and 8-6). Trees severely affected by tatters 

have thin crowns in spring (fig. 8-7). A new flush of undamaged leaves usually develops, 

although some twig and branch dieback may be present.

Cause

Leaf tatter has been associated with several environmental factors, although causation 

in many cases has not been shown. Frost or freeze that occurs in spring as buds are beginning to open may be a potential cause for some cases. Injury is thought to be initiated 

when bud scales loosen and tiny areas of tender 

leaf tissue become exposed to cold temperatures. 

As leaves unfold and expand, the injured or 

killed areas develop into holes (fig. 8-1).

Wind tunnel experiments have produced 

tatterlike symptoms in maple, and tattering is 

frequently observed on young, tender foliage 

of many trees following strong winds in spring. 

One study of sugar maple leaves suggests that 

the anatomical makeup of the leaves of some 

cultivars may play a role in their susceptibility to 

tatter.

Figure 8-7—Thin crowns of hackberry caused 

by leaf tatters (Laurie J. Stepanek, Nebraska 

Forest Service, used with permission).
40 USDA Forest Service RMRS-GTR-335. 2016.

Leaf tatters of oak and hackberry has been shown in studies to be caused by herbicides containing acetochlor, metolachlor, and dimethenamid. These herbicides are 

frequently used in corn production, which could explain the high number of tatters 

reports occurring in the Midwest, where corn production is common.

Leaf tatters symptoms develop when herbicide exposure occurs as the leaves are unfolding and expanding. Leaves still in the bud and leaves fully expanded are not affected. 

Exposure may potentially occur through spray drift, volatilization, or contaminated rain 

or dust, thereby affecting trees both near and far from the application.

Damage

Leaf tatter and leaf tatters typically affect only the appearance of the trees. Severely 

affected trees may produce a new flush of leaves if they have sufficient energy reserves, 

but the stress of leaf loss can make trees more susceptible to other damaging agents. 

Branch dieback or tree death may occur in trees that have been severely affected over 

several years, especially if they have suffered prior stress, such as from drought or other 

poor growing conditions.

Management

Sugar maple selections less susceptible to maple tatter include Legacy® Maple 

(Acer saccharum ‘Legacy’, PP4,979) Commemoration® Maple (Acer saccharum

’Commemoration’, PP5,079) Oregon Trail® Maple (Acer saccharum ‘Hiawatha1’) John 

Pair Maple, (Acer saccharum ‘John Pair’) Autumn Splendor Maple (Acer saccharum

‘Autumn Splendor’) Flashfire® Maple (Acer saccharum ‘JFS-Caddo2’, PP23,361) 

‘Oregon Trail’ is a northeastern Kansas selection; ‘Fall Fiesta’ originated in Minnesota; 

and the last three are Caddo maples, which are a population of sugar maple in western 

Oklahoma.

Oak and hackberry tatters may be reduced by application of crop herbicides before or 

well after budbreak to avoid sensitive stages of leaf growth. Large droplet size, low boom 

heights, and the addition of drift inhibitors can help minimize drift. Soil incorporation can 

minimize volatilization.

Selected References

Conley, M.E.; Paparozzi, E.T.; Pair, J.C.; Stroup, W.W. 1995. Leaf tatter in Acer 

saccharum: an anatomical link. International Journal of Plant Science. 156(3): 

303–310.

Flickinger, A. 2007. Iowa forest health. 56 p. http://iowadnr.gov/Environment/Forestry/

ForestHealth.aspx. (November 27, 2013).

Miller, A. 2010. Chloroacetamide spray drift and leaf tatters in hackberry. Lincoln, 

NE: University of Nebraska. 91 p. M.S. thesis. http://digitalcommons.unl.edu/

natresdiss/12/. (November 27, 2013).

Samtani, J.B.; Masiunas, J.B.; Appleby, J.E. 2008. Injury on white oak seedlings from 

herbicide exposure simulating drift. HortScience. 43(7): 2076–2080.

Samtani, J.B.; Masiunas, J.B.; Appleby, J.E. 2010. White oak and northern red oak leaf 

injury from exposure to chloroacetanilide herbicides. HortScience. 45(4): 696–700.

Wilson, J. 1980. Macroscopic features of wind damage to leaves of Acer pseudoplatanus

L. and its relationship with season, leaf age, and windspeed. Annals of Botany. 46: 

303–311.
USDA Forest Service RMRS-GTR-335. 2016. 41

9. Marssonina Leaf Spot and Blight of Populus

Species

Michael E. Ostry and Jared M. LeBoldus; revised from John E. 

Watkins and David S. Wysong (Riffle and Peterson 1986)

Marssonina leaf spot, a widespread and serious disease of native and hybrid poplars 

(Populus spp.), can severely defoliate susceptible genotypes well before normal leaf drop. 

This disease is caused by at least four different species of fungi in the genus Marssonina. 

Hosts and Distribution

There are four relatively well studied species of Marssonina affecting poplar in 

Europe, North America, Asia, Australia, and New Zealand: M. brunnea (teleomorph = 

Drepanopeziza punctiformis), M. castagnei (D. populi-albae), M. populi (D. populorum), 

and M. balsamiferae (teleomorph unknown). These fungi affect balsam poplar 

(P. balsamifera), plains cottonwood (P. deltoides subsp. monilifera), quaking aspen 

(P. tremuloides), white poplar (P. alba), and various hybrid poplars. Marssonina species 

are known to be host specific. On willows (Salix spp.), M. brunnea and M. salicicola

(teleomorph = Drepanopeziza sphaerioides) are the major species of importance. A fifth 

taxon, M. populicola, is known to affect poplars in China. Species in section Populus

(aspens, white poplars) are susceptible to M. brunnea f. sp. trepidae, and species in section Aigeiros (cottonwoods, black poplars) are susceptible to M. brunnea f. sp. brunnea. 

M. balsamiferae was reported occurring on P. balsamifera in Ontario and Manitoba. 

M. castagnei infects P. alba, and M. populi is common on P. tremuloides. Marssonina 

populicola has been reported in China on P. davidiana, P. euphratica, P. euramericana, 

P. laurifolia, P. pseudosimonii, P. simonii, and P. tomentosa. Severe defoliation of 

Populus species and their hybrids by Marssonina has been reported across the Great 

Plains.

Symptoms and Signs

Dark brown flecks (fig. 9-1), often with yellow margins (fig. 9-2), appear on leaves 

within a few weeks of leaf emergence. These spots gradually enlarge to 1 to 2 mm in 

diameter. Individual spots on severely affected leaves may coalesce to form angular, 

necrotic blotches. Diseased leaves on affected trees appear smaller than normal, turn 

yellow to bronze, and fall prematurely (fig. 9-3). Symptoms develop progressively upward into the crown. If viewed from a distance, the diseased leaves appear bronzed. On 

highly susceptible clones lens-shaped lesions develop on petioles (fig. 9-4) and currentyear shoots (fig. 9-5). Subcuticular fruiting bodies (acervuli) containing asexual spores 

(conidia) measuring 11 to 16 μm × 3.5 to 7.0 μm (fig. 9-6) in a white gelatinous mass 

form within leaf, petiole, and stem lesions of infected hosts.

Figure 9-1—Angular leaf spots caused 

by Marssonina brunnea on a poplar leaf 

(Michael E. Ostry, U.S. Forest Service).

Figure 9-2—Marssonina leaf spot on aspen with 

necrotic leaf spots with yellow margins (Joseph 

G. O’Brien, U.S. Forest Service).
42 USDA Forest Service RMRS-GTR-335. 2016.

Figure 9-4—Elliptical lesions on petioles of poplar leaves caused by Marssonina

brunnea (Michael E. Ostry, U.S. Forest Service).

Figure 9-5—Lesions on poplar stems caused by Marssonina brunnea (Michael E. 

Ostry, U.S. Forest Service). 

Figure 9-3—Healthy green poplar leaf and yellow poplar leaf infected by 

Marssonina brunnea (Michael E. Ostry, U.S. Forest Service). 
USDA Forest Service RMRS-GTR-335. 2016. 43

Disease Cycle

The fungus overwinters in fallen leaves and in infected shoots. In spring, ascospores 

produced in the leaves and conidia from lesions on the shoots are released in wet weather. 

Leaves and new shoots are infected throughout the growing season by rain-splashed 

conidia, intensifying disease severity. Epidemics coincide with extended or frequent wet 

weather.

Damage

In nurseries and plantations, Marssonina leaf spot causes only slight damage to cuttings and young seedlings until sufficient inoculum develops on infected shoots and on 

fallen leaf debris. In more established plantings and in native stands, repeated outbreaks 

result in branch and twig dieback and predispose trees to other pathogens or pests and to 

injury from low temperatures. 

Management

The preferred method of control is to plant clones resistant to the disease. Pruning diseased twigs and branches and removal of fallen leaves may also limit disease incidence 

and severity. Fungicide has also proven to be an effective means of control; however, 

there is currently no fungicide labeled for use in plantations. To avoid spreading this 

pathogen to new areas, cuttings should only be taken from disease-free shoots. However, 

it is important to remember that infected leaves and shoots may remain asymptomatic and 

that this pathogen is seed borne. As a result, care must be given to avoid inadvertent shipment of infested seed or asymptomatic plant material. 

Selected References

Newcombe, G. and Callan, B.E. 1988. First report of Marssonina brunnea f. sp. brunnea

on hybrid poplar in the Pacific Northwest. Plant Disease. 81: 231.

Spiers, A.G. 1983. Host range and pathogenicity studies of Marssonina brunnea to 

poplars. European Journal of Forest Pathology. 13: 181–196.

Spiers, A.G. 1984. Comparative studies of host specificity and symptoms exhibited by 

poplars infected with Marssonina brunnea, Marssonina castagnei, and Marssonina 

populi. European Journal of Forest Pathology. 14: 202–218.

Spiers, A.G. 1988. Comparative studies of type and herbarium specimens of Marssonina 

species pathogenic to poplars. European Journal of Forest Pathology. 18: 140–156.

Spiers, A.G. 1989. Introduction of poplar and willow pathogens into New Zealand and 

their effects. New Zealand Journal of Forestry Science. 19: 347–352.

Spiers, A.G. 1998. Melampsora and Marssonina pathogens of poplars and willows in 

New Zealand. European Journal of Forest Pathology. 28: 233–240.

Spiers, A.G.; Hopcroft, D.H. 1998. Morphology of Drepanopeziza species pathogenic to 

poplars. Mycological Research. 102: 1025–1037.

Figure 9-6—Conidia of Marssonina brunnea. Scale bar is 10 μm 

(Michael E. Ostry, U.S. Forest Service).
44 USDA Forest Service RMRS-GTR-335. 2016.

10. Melampsora Leaf Rust of Cottonwood, Aspen, and 

Willow

Phillip A. Mason and Jared M. LeBoldus; revised from Glenn W. 

Peterson and Robert W. Stack (Riffle and Peterson 1986)

Cottonwood (Populus deltoides), aspen (Populus spp.), and willow (Salix spp.), 

including both native species and their hybrids, are susceptible to infection by the fungal 

leaf rust pathogens in the genus Melampsora. On highly susceptible trees the disease 

results in premature leaf drop, which can be deleterious to tree health and vigor. 

Hosts and Distribution

Melampsora leaf rust is widely distributed, affecting numerous Populus species, 

hybrids, and clones from North Dakota to Texas. In the Great Plains the Melampsora

pathogen is often found on various Populus species and their hybrids, quaking aspen 

(P. tremuloides), and willow (Salix spp.). The alternate host of these rust species is 

larch (Larix spp.). The most common rust species in the Great Plains are Melampsora 

medusae on cottonwood and aspen and Melampsora epitea on willow. However, other 

Melampsora species have been reported to be expanding their known host ranges, and 

natural rust hybrids are occurring.

Symptoms and Signs

The most commonly recognized sign of this disease is orange pustules (uredinia) on 

the lower (abaxial) leaf surface (figs. 10-1 and 10-2). The orange powdery pustules are 

composed of a mass of urediniospores that usually form on the underside of leaves in the 

lower crown of trees. As the leaves become severely diseased, they begin to senesce. At 

this time brown to black crusts (telia) begin to form on leaves among the orange uredinia 

and become prevalent on fallen leaves. Premature leaf drop occurs in midsummer.

Disease Cycle

Melampsora medusae and M. epitea require two hosts to complete their life cycle. The 

fungi cycle most often between the genera Populus and Larix. In spring the disease cycle 

begins on fallen, infected Populus leaves, when the overwintering teliospores on telia 

germinate, giving rise to basidia, another 

spore-bearing structure where basidiospores form. The basidiospores can infect 

only the conifer host (larch), which later 

gives rise to aecia, a spore-bearing structure on larch needles where aeciospores 

develop. Aeciospores are windblown 

and infect the Populus or Salix host (cottonwood, aspen, or willow). Uredinia then 

develop and production of the urediniospores occurs. These spores are released 

and infect other poplar or willow hosts 

(figs. 10-1 and 10-2) and often re-infect 

the same host plant continuously until leaf 

drop. Telia form on the leaves in late summer, and the infected, fallen leaves serve 

as the overwintering site for the pathogen, 

completing the complex life cycle of 

the fungus. In many areas of the Great 

Plains where the larch is not present, it is 

believed that the urediniospores are spread 

via wind in a north to south direction. Figure 10-1—Orange powdery urediniospores 

of Melampsora on the abaxial surface of willow 

leaves (Phillip A. Mason, Animal and Plant 

Health Inspection Service).
USDA Forest Service RMRS-GTR-335. 2016. 45

Figure 10-2—Orange powdery urediniospores of Melampsora on the abaxial surface 

of a hybrid poplar leaf (Kelsey L. Dunnell, North Dakota State University, used with 

permission). 

Damage

Depending on leaf rust severity, premature leaf drop can occur, resulting in loss of 

vigor and reducing the tree’s ability to defend against other stresses. Premature defoliation in successive years can reduce carbohydrate reserves and contribute to the decline of 

Populus species in the Great Plains. Although M. epitea has been reported to cause stem 

cankers on young willow stems, cankers have not been reported to occur with M. medusae on cottonwood or aspen. 

Management

Fungicides are effective in protecting trees from infection by Melampsora, but this 

is not an economically practical solution when control is desired in larger plantings like 

that of a windbreak. The disease impact in the Great Plains can be reduced by planting 

a variety of Populus hybrids and clones, specifically those that exhibit various levels 

of resistance to Melampsora leaf rust. The recognition of Eurasian poplar leaf rust 

(Melampsora × columbiana), a new Melampsora hybrid in the Pacific Northwest, and M. 

occidentalis in the central United States suggests that a Populus hybrid or clone resistant 

to one species of Melampsora can be susceptible to another Melampsora species. Use of 

trees with various levels of resistance to Melampsora species should help limit the impact 

of the disease on larger plantings of Populus. 

Selected References

McCracken, A.R.; Dawson, M. 2003. Rust disease (Melampsora epitea) of willow (Salix 

spp.) grown as short rotation coppice (SRC) in inter- and intra-species mixtures. 

Annals of Applied Biology. 143: 381–393.

Moltzan, B.D.; Stack, R.W.; Mason, P.A.; Ostry, M.E. 1993. First report of Melampsora 

occidentalis on Populus trichocarpa in the central United States. Plant Disease. 77: 

953.

Newcombe, G.; Stirling, B.; McDonald, S.; Bradshaw, S. 2000. Melampsora × 

columbiana, a natural hybrid of M. medusae and M. occidentalis. Mycological 

Research. 104: 261–274.

Ostry, M.E.; Anderson, N.A. 2001. Melampsora leaf rust of willow causing a stem canker 

in Minnesota. Plant Disease. 85: 229.

Ziller, W.G. 1965. Studies of western tree rusts. VI. The aecial host ranges of

Melampsora albertensis, M. medusae and M. occidentalis. Canadian Journal of 

Botany. 43: 217–230.
46 USDA Forest Service RMRS-GTR-335. 2016.

11. Mycosphaerella Leaf Spot of Ash

Judith O’Mara and Megan M. Kennelly

Mycosphaerella leaf spot of ash is caused by two species of Mycosphaerella

(M. effigurata and M. fraxinicola). The disease occurs on all ash (Fraxinus) species 

grown throughout the Great Plains. In most years the disease occurs late in the growing 

season and causes minor leaf spotting and defoliation. 

Hosts and Distribution

White ash (F. americana) and green ash (F. pennsylvanica) are the dominant ash 

species in the Great Plains. The composition of ash in urban and woodland plantings 

within the Great Plains ranges from 10 to 15 percent in Kansas, Oklahoma, and parts of 

Missouri to greater than 50 percent in North and South Dakota. The Mycosphaerella leaf 

spot disease complex occurs primarily on ash in the eastern states of the Great Plains. No 

resistant ash species are known, although they can differ in their level of susceptibility. 

Symptoms and Signs

The symptoms for both Mycosphaerella species are distinctive, but can be difficult to 

distinguish visually as both pathogens may be present at the same time. M. fraxinicola

produces large, irregular leaf spots that start out light green and then turn brown with 

yellow halos (5 to 15 mm) (fig. 11-1). Individual lesions may coalesce to form larger 

necrotic blotches. M. effigurata produces numerous small, yellow to brown spots (1 to 

3 mm) on the upper leaf surface. At midseason, masses of white asexual spores 

(conidia) may be found in acervuli on 

the upper leaf surface (fig. 11-2). Late in 

the fall, black spots (stromata) develop 

in lesions on the lower leaf surface. The 

disease starts on the lowest portion of the 

tree and moves up (fig. 11-3). Severely 

infected trees may prematurely defoliate 

four to six weeks early.

Disease Cycle

Mycosphaerella leaf spot of ash 

is caused by two pleomorphic fungi, 

M. effigurata and M. fraxinicola. M. 

effigurata initiates symptoms earlier in 

summer; M. fraxinicola tends to occur August through 

September. During dry years, the impact of the disease 

is minor. During wet growing seasons in Kansas, M. 

fraxinicola triggers visible symptoms by early-to-mid 

July and entire trees may be completely defoliated. 

M. fraxinicola has a complex life cycle with three 

stages. Pseudothecia overwinter and mature in ash leaf 

litter. The ascospores are 8 to 10 μm × 4 to 5 μm, oval, 

two-celled, and colorless. Initial infection by ascospores 

takes place late spring to early summer. The anamorph 

(Pseudocercosporella fraxini; syn. Cercospora fraxini 

and Cylindrosporium fraxini) develops in summer and 

produces masses of colorless, cylindrical, multicelled 

conidia primarily on the upper leaf surface. The spermatial state of the Asteromella type produces tiny black 

structures that appear late summer in lesions on the 

lower leaf surface. 

Figure 11-1—Mycosphaerella leaf spot on ash 

(Ward Upham, Kansas State University, used with 

permission).

Figure 11-2—Mycosphaerella 

leaf spot with white masses 

of conidia produced by 

Pseudocercosporella fraxini 

(Megan M. Kennelly, Kansas 

State University, used with 

permission).
USDA Forest Service RMRS-GTR-335. 2016. 47

M. effigurata can also be found on ash 

leaves infected by M. fraxinicola. Lesions 

of M. effigurata tend to be smaller and 

more numerous. Ascospores are 4 to 14 μm 

long, overwinter in leaf litter, and initiate 

infection in late spring to early summer. 

During midsummer, acervuli produce clear, 

two-celled conidia on the underside of the 

leaf (Septoria fraxinicola; syn. Marssonina 

fraxini). Late in the season a black 

Asteromella spermagonial state follows on 

the underside of the leaf.

Damage

Mycosphaerella leaf spot can be found 

on ash trees during any given year. In most 

years, it is a late-season disease that causes 

minor shedding. During wet summers, 

high disease pressure can result in rapid 

defoliation and complete leaf loss one to 

two months earlier than normal autumn 

leaf fall. Leaf spot diseases that occur late 

in the growing season are generally considered to have minimal impact on tree health. 

However, successive years of wet weather 

and high disease pressure can result in ash 

trees with poor emergence, low vigor, and 

branch dieback. 

Ash trees of all ages are susceptible to Mycosphaerella leaf spot, with younger trees in 

nursery settings more severely affected. Owing to the continuing spread of the emerald 

ash borer (Agrilus planipennis), ash trees are no longer widely grown in nurseries. Thus, 

until a time when effective controls for EAB or resistant species or genotypes are developed, the impact of Mycosphaerella leaf spot of ash on young trees will be primarily in 

landscaped and wooded areas.

Management

Mycosphaerella leaf spot of ash generally occurs late in the growing season with 

minimal defoliation preceding normal fall maturity. When wet years result in extensive 

defoliation, raking leaf litter in the fall may help reduce overwintering disease inoculum. 

Trees undergoing multiple years of heavy disease pressure will benefit from proper pruning to promote good air flow and regular irrigation to help maintain tree vigor. Chemical 

control measures are not available for this disease complex. 

Selected References

Connors, I.L. 1967. An annotated index of plant diseases in Canada. Agric. Publ. 1251. 

Ottawa, ON: Canada Department of Agriculture. 381 p. 

Hepting, G.H. 1971. Diseases of forest and shade trees of the United States. Agric. 

Handb. 386. Washington, DC: U.S. Department of Agriculture, Forest Service. 658 p. 

Sinclair, W.; Lyon, H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: Cornell 

University Press. 660 p. 

Wolf, F.A. 1939. Leaf spot of ash and Phyllosticta viridis. Mycologia. 31: 258–266. 

Wolf, F.A.; Davidson, R.W. 1941. Life cycle of Piggotia fraxini, causing leaf disease of 

ash. Mycologia. 33: 526–539.

Figure 11-3—Extensive defoliation on ash resulting from Mycosphaerella fraxinicola (Ward 

Upham, Kansas State University, used with 

permission).
48 USDA Forest Service RMRS-GTR-335. 2016.

12. Powdery Mildew of Lilac

John Ball; revised from Richard Dorset and Michael W. Ferguson 

(Riffle and Peterson 1986)

Lilacs (Syringa spp.) are some of the most common shrubs planted in the Great Plains, 

and the many species within this genus can be found in ornamental plantings as well as 

in windbreaks. Common lilac (S. vulgaris) is the most common lilac species in the region 

and is prized as much for its winter hardiness and adaptation to the harsh growing conditions of the Great Plains as for its fragrant late-spring flowers.

Lilacs are not without their stressors, however, and lilac/ash borer (Podosesia syringae), bacterial blight, and powdery mildew are all frequent problems. Powdery mildew 

is a disease often noticed by people in late summer or early fall as they pass by infected 

plants.

Hosts and Distribution

Powdery mildew is the common name for a disease caused by a number of closely 

related fungi that produce similar symptoms in their hosts. There are almost 100 different powdery mildew species affecting more than 10 different genera of woody plants 

in North America alone. Woody plant genera most commonly infected with powdery 

mildew are Aesculus (buckeyes), Quercus (oaks), Rosa (roses), Vitis (grapes), and lilacs. 

A powdery mildew species will not spread among these different genera, as mildew fungi 

are specific to a host genus. Although powdery mildew is common on many tree species, 

the disease is primarily a concern on lilacs. The remainder of this chapter will focus on 

powdery mildew of lilacs (Erysiphe syringae). This is a different disease from downy 

mildew, a disease that rarely affects lilacs.

Symptoms and Signs

Symptoms appear first on the lower leaves and spread upward along the branch as the 

growing season progresses. The upper surface of infected leaves develops small white to 

gray, almost dusty-appearing, patches of the mycelium (fig. 12-1). These colonies continue to enlarge through the summer and by early fall the entire leaf, and most of the leaves 

on the plant, may be covered with a white powdery material. Although the leaf surface 

appears powdery, the foliage itself may turn yellow. Severely infected leaves also become 

distorted and twisted and even occasionally wilt. The wilting is sometimes mistaken for 

injury from herbicide drift.

Figure 12-1—Late-summer appearance of common lilac infected with powdery 

mildew (John Ball, South Dakota State University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 49

Disease Cycle

Small, almost pinpoint-size brown to black structures (ascocarps) begin development in late summer within the colonies and are easily seen with a 10× hand lens. The 

pathogen overwinters in the fallen, infected leaves as partially developed ascospores in an 

ascus enclosed in a fruiting body (chasmothecium; fig. 12-2). The ascospores mature during the early spring. As daytime air temperatures exceed 70 °F, they are released through 

cracks in the fruiting body and are carried by the wind to susceptible hosts. This is also 

the time when lilac shoots and foliage begin to expand. Once spores land on the leaf, germination can occur within a few hours if environmental conditions are favorable. Unlike 

many other pathogens, free water inhibits germination of the spores. The disease requires 

different environmental conditions for spore formation and dispersal. The disease develops best with repeated cycles of cool (60 °F), humid nights (spore formation) and warm 

(80 °F), dry days (spore dispersal). Infection is generally limited to the succulent new 

growth; mature leaves are resistant. Ascospores initiate the disease cycle on a plant, with 

secondary infection by conidia during the growing season. The conidia are invisible to 

the naked eye, but chains of conidia on the hyphae (conidiophores) contribute to the overall powdery appearance. The conidiophores generally arise from the epiphytic hyphae.

Damage

The disease appears most commonly on the foliage, though occasionally shoots can 

be affected. Powdery mildew is unusual in that it lives on top of as well as beneath the 

leaf surface. The hyphae live on the leaf surface but also produce haustoria that penetrate 

the epidermis and extend into the elongated palisade mesophyll cells that are the primary 

sites for photosynthesis. The haustoria extract water and nutrients from these cells. The 

disease is rarely fatal to lilacs. The appearance of the symptoms is usually the major concern with the lilac owner, who may view them as aesthetically unattractive.

Management

The simplest management of powdery mildew is to alter the growing environment, 

making it less favorable for development of the disease. This approach requires pruning 

and clearing lilac beds to decrease night humidity and improve air flow. Removing overhanging trees or, if the tree cannot be removed, pruning off the lower limbs (those lower 

than 12 feet) may provide sufficient air flow and reduced night humidity to minimize 

Figure 12-2—Closeup of overwintering fruiting body (chasmothecium) and appendages of powdery mildew species (Harold Larsen, Colorado State University, 

used with permission).
50 USDA Forest Service RMRS-GTR-335. 2016.

infection. However some lilacs, particularly the common lilac and its many cultivars, may 

be covered with powdery mildew even in open growing conditions. 

Another management option is to plant lilac species that are rarely affected by powdery mildew. Not all of these species have the same form or flowering characteristics of 

the common lilac but may be acceptable substitutes in many planting locations.

The lilac species and hybrids in which powdery mildew is rarely a problem in the Great 

Plains include: early flowering lilac (S. × hyacinthiflora), Meyer lilac (S. meyeri), littleleaf lilac (S. microphylla), Manchurian lilac (S. patula), Pekin tree lilac (S. pekinensis), 

Preston lilac (S. × prestoniae), Japanese tree lilac (S. reticulata), late lilac (S. villosa).

Chinese lilac (S. × chinensis) can occasionally be infected with powdery mildew 

and Persian lilac (S. × persica) and its cultivars may also become infected, but the 

species most severely infected is the one that is planted the most: the common lilac. 

Almost all cultivars of common lilac are susceptible to this disease, including the French 

hybrids; however, there are a few cultivars that may not become infected or at least 

exhibit barely noticeable symptoms of the disease: ‘Adelaide Dunbar,’ ‘Charles Joly,’ 

‘Hulda,’‘Jan Van Tol,’ and ‘Katherine Havemeyer.’

If the disease does not appear until late summer, there is little concern that the disease 

will seriously harm the health of the lilac. The shoot growth and flowering will not be reduced the next spring. If the disease begins to develop earlier in the season, the infection 

may result in reduced growth. Regardless of whether the disease symptoms appear early 

or late in the growing season, the appearance of “dusty” leaves may be unacceptable to 

the plant’s owner. 

Because the disease-causing fungus occurs primarily on the leaf surface, powdery 

mildew is much easier to manage with fungicides than are many other diseases. The 

choice of fungicides will vary with availability and registration within a particular state 

or province, but generally a fungicide labeled for powdery mildew on lilacs can be found 

in garden stores. A fungicide application can be made when the disease symptoms are 

first noticed and then continued every two weeks while conditions are favorable for the 

disease. Once lilac finishes its growth, usually in midsummer, leaves become resistant to 

the disease and treatments can be discontinued.

Selected References

Clement, D.L.; Gill, S.A.; Potts, W. 1994. Alternatives for powdery mildew control on 

lilac. Journal of Arboriculture. 20: 227–230.

Hibben, C.R.; Walker, J.T.; Allison, J.R. 1977. Powdery mildew rating of lilac species 

and cultivars. Plant Disease Reporter. 55: 475–478.

Mmbaga, M.T.; Suave, R.J.; Nnodu, E.; Zhou, S. 2005. Multiple disease resistance to 

powdery mildew, bacterial blight and alternaria blight in lilacs (Syringa spp). Journal 

of Arboriculture. 31: 1–9.

Schnathorst, W.C. 1965. Environmental relationships in the powdery mildews. Annual 

Review of Phytopathology. 3: 343–366.
USDA Forest Service RMRS-GTR-335. 2016. 51

13. Tar Spots of Maple

Aaron D. Bergdahl and Kelsey L. Dunnell

The fungi that cause tar spots of maple are becoming a concern in the Great Plains as 

the popularity of planting maple (Acer) species and their cultivars increases throughout 

the region. Although tar spots do not often severely affect tree health, they have the potential to significantly reduce the aesthetic appeal of trees. 

Hosts and Distribution

Tar spot is a disease of maple leaves caused by one of three species of fungi in the genus Rhytisma. All three fungi occur in the central United States. Tar spot pathogens have 

been known to cause diseases on maple species throughout the eastern and Great Lakes 

regions, in the Pacific Northwest, and in Manitoba, Canada. In the central United States, 

tar spots have been observed on Acer species in Kansas, Missouri, Nebraska, North 

Dakota, South Dakota, and Wyoming. Refer to table 13-1 for maple hosts and associated 

Rhytisma species. 

Symptoms and Signs

In mid- to late July one or more small greenish-yellow spots appear on the upper leaf 

surface indicating early infection (fig. 13-1a). In late summer conspicuous black, tarlike 

structures form within the chlorotic areas (fig. 13-1b,c,d). The lower leaf surface opposite 

the tarlike structures is cupped and less black. 

The tar spot fungus most often associated with Norway maple (A. platanoides) is 

caused by R. acerinum (fig. 13-1b) and was apparently introduced from Europe in the 

early 1900s and first formally reported in North America in 1942. Tar spots caused by

R. acerinum can exceed 0.5 inch in diameter, which is why it has been referred to as 

“giant tar spot” in some earlier references. In the Great Plains the fungus R. americanum 

(fig. 13-1c) primarily infects red maple (A. rubrum), silver maple (A. sacharrinum), and 

their hybrids. In the past, the causal fungus R. americanum has often been misidentified 

as R. acerinum and the confusion between these two fungal pathogens continues owing 

to their similar appearance and previously used common names. The extent to which 

speckled tar spot (R. punctatum) (fig. 13-1d) occurs in the Great Plains is uncertain. Other 

Rhytisma species commonly cause tar spot on willow (Salix spp.), holly (Ilex spp.), and 

tuliptree (Liriodendron tulipifera), although these are not discussed here.

Table 13-1—Maple species and their susceptibility to three species of tar spot fungi.

Tar spot species

Maple species Rhytisma acerinum R. americanum R. punctatum

Acer campestre (hedge maple) X

A. circinatum (vine maple) X X

A. ginnala (Amur maple) X

A. glabrum (Rocky Mountain maple) X X

A. negundo (boxelder) X X X

A. macrophyllum (bigleaf maple) X X

A. pensylvanicum (striped maple) X

A. platanoides (Norway maple) X

A. pseudoplatanus (sycamore maple) X X X

A. rubrum (red maple) X X X

A. saccharinum (silver maple) X X X

A. saccharum (sugar maple) X X X

A. spicatum (mountain maple) X X X 

Adapted from Hsiang and Tian (2007).
52 USDA Forest Service RMRS-GTR-335. 2016.

Disease Cycle

Ascospores of the tar spot fungus generally begin to develop in the black spots 

(stromata) before leaves are cast in fall and continue development when leaves are on 

the ground. In spring the stroma begins to split, and mature needle-shaped ascospores 

are forcibly ejected upward from apothecia on fallen leaves during favorable conditions. 

The spores are ejected into the air for a limited distance and then carried upward on air 

currents, where they make contact with expanding foliage of susceptible trees. Germ 

tubes from the germinating spores enter the leaves through stomata, and a black gummy 

substance is produced within the tissues.

Symptoms are generally visible within two months of infection. At this time each 

lesion appears as a defined blackened area fringed with a yellowish-green border. First, 

numerous spore-producing structures develop in a group at the center of the stroma. They 

resemble small pimples with a minute hole in the center, and represent the imperfect 

(asexual) stage. Simple or branched conidiophores are formed within each fruiting body, 

producing clear (hyaline) or partially clear (subhyaline), rodshaped spores that measure 

6 μm × 1 μm. It has been proposed that these single-celled spores are not infectious 

and play a role in sexual reproduction (therefore, they are often referred to as spermatia 

formed in spermagonia), although this has not been confirmed. Later, apothecia begin to 

Figure 13-1—Symptoms of infection: (A) Typical symptoms of early infection by Rhytisma acerinum (George W. Hudler, Cornell University, used with permission); (B) Late-season tar spot (R. 

acerinum) on Norway maple (Aaron D. Bergdahl, North Dakota Forest Service, used with permission); (C) R. americanum on silver maple (Simeon Wright, Missouri Department of Conservation, 

used with permission); (D) Infection by speckled tar spot R. punctatum (George W. Hudler, 

Cornell University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 53

develop within this tissue before 

the leaves drop. Stromata continue 

to expand, developing radiating 

wrinkles in which apothecia 

form (fig. 13-2a). Asci slowly 

develop through the winter and by 

early spring their development is 

completed. Each ascus within an 

apothecium (fig. 13-2b) contains 

eight needlelike ascospores (fig. 

13-2c), approximately 130 μm × 

10 μm. The stroma splits along the 

radiating wrinkles and ascospores 

are ejected, completing the disease 

cycle (fig. 13-3).

Damage

Tar spot seldom severely affects the health of trees and is not 

known to kill established trees, 

although it may lead to seedling 

mortality after heavy infection. In 

sheltered situations, such as in the 

forest, trees can become heavily 

infected and defoliate prematurely, 

whereas open-grown trees are 

typically less infected. However, 

during years of prime conditions 

for infection, large open-grown 

trees can have a severe level of infection by R. acerinum. Successive 

years of defoliation may constitute 

a significant stress, predisposing 

trees to other pests. However, two 

successive years of significant 

premature defoliation seldom 

occurs because heavily infected 

leaves are prematurely shed. Thus, 

the stroma on shed leaves has not 

had sufficient time to fully develop 

asci for reinfection the next spring. 

Further, in a forest setting early shed leaves are often covered by leaves of other tree species, creating a barrier to spore dispersal. Premature defoliation of infected leaves gives 

the appearance of crown thinning.

Management

The disease can be controlled in shade trees by raking and burning fallen leaves to destroy overwintering inoculum. Efficient composting of leaves is also an effective practice 

for reducing inoculum. Typically, use of fungicides to control this disease is discouraged. 

If spraying is warranted, copper-based fungicides are considered satisfactory. Although 

management of R. acerinum may be achieved with one well-timed fungicide application, 

just as the leaves on Norway maple reach full expansion, applying fungicide at two- to 

three-week intervals during leaf emergence may provide best results. 

Figure 13-2—Stages of disease cycle of tar spot of 

maple disease: (A) Splitting Rhytisma stroma after an 

overwintering period prior to spore dispersal; (B) Crosssection of mature Rhytisma apothecium; (C) Asci and 

ascospores of Rhytisma species (George W. Hudler, 

Cornell University, used with permission).

B

C

A
54 USDA Forest Service RMRS-GTR-335. 2016.

Selected References

Hudler, G.W.; Banik, M.T.; Miller, S.G. 1987. Unusual epidemic of tar spot on Norway 

maple in upstate New York. Plant Disease. 71: 65–68. 

Hudler, G.W.; Jensen-Tracy, S.; Banik, M.T. 1998. Rhytisma americanum sp. nov: a 

previously undescribed species of Rhytisma on maples (Acer spp.). Mycotaxon. 68: 

405–416. 

Hsiang, T.; Tian, X.L. 2007. Sporulation and identity of tar spot of maple in Canada. Acta 

Silvatica & Lignaria Hungary. Special Edition: 71–74.

Lapointe, M; Brisson, J. 2011. Tar spot disease on Norway maple in North America: 

quantifying the impacts of a reunion between an invasive tree species and its adventive 

natural enemy in an urban forest. Ecoscience 18(1): 63–69.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Figure 13-3—Disease cycle of tar spot of maple (David Haasser, North Dakota State University,

used with permission, Adapted from Riffle and Peterson, 1986).
USDA Forest Service RMRS-GTR-335. 2016. 55

14. X-Disease of Chokecherry

James A. Walla; revised from Glenn W. Peterson and David W. 

Johnson (Riffle and Peterson 1986)

Chokecherry (Prunus virginiana) is a native shrub used in conservation, ornamental, 

and fruit production plantings in the central and northern Great Plains. Chokecherry in 

many of these plantings is infected by X-disease phytoplasmas, ‘Candidatus Phytoplasma 

pruni,’ RFLP group 16sRIII-A. Phytoplasmas are closely related to Gram-positive 

bacteria. Previous names for X-disease include virus-X, eastern X-disease, and western 

X-disease.

Hosts and Distribution

X-disease seriously damages common chokecherry (Prunus virginiana L. var. 

virginiana), sweet (P. avium) and sour (P. cerasus) cherries, and peach (P. persica). Also 

susceptible are western chokecherry (P. virginiana L. var. demissa (Nutt.) Torr.), apricot 

(P. armeniaca), domestic plum (P. americana), Japanese plum (P. salicina), and some 

other Prunus species. The red-leaf strains of common chokecherry, such as ‘Schubert’ 

chokecherry and ‘Canada Red’ cherry, are also damaged. Leafhoppers are thought to be 

the primary vector, but the specific vectors are not known in the Great Plains. X-disease 

occurs in the northern half of the United States and southern half of Canada, including the 

central and northern Great Plains states and the Prairie Provinces.

Symptoms

X-disease usually causes a slow decline, but symptoms vary with phytoplasma genotype, host species and genotype, disease development stage, host age, and site conditions. 

In chokecherry, one or more years after infection, leaves on an infected stem typically 

become greenish yellow a few weeks before normal onset of fall color. They may have 

green tissue remaining along the veins and an orange or red tinge or necrosis on their borders, and there may be leaves with different stages of disease on a single shoot (fig. 14-1). 

Leaf discoloration, deformation, and stunting begin earlier and become more pronounced 

each subsequent year, until first discoloration is at budbreak over the entire plant and 

the leaves may turn deep red or orange by six to eight weeks after budbreak (fig. 14-2). 

Moderately or severely diseased leaves feel thicker and waxier than healthy leaves. 

Shoots and roots are stunted, and rosettes may result from shortened internodes at shoot 

tips. Fruit may be small and somewhat pointed and may ripen to bright red, not the normal dark purple of healthy fruit (fig. 14-3a). Buds growing from current-season leaf axils 

and small late-season flowers may develop (fig. 14-3b). Twigs do not harden off as early, 

and twig dieback may result. After two or more years, branch, whole stem, or whole plant 

dieback may occur. In red-leaf chokecherry strains, X-disease causes similar decline, but 

leaf discoloration is masked; severely diseased leaves are lighter red than healthy leaves.

Figure 14-1—Chokecherry leaves with X-disease symptoms. Leaves become various shades of 

(A) yellow, (B) red, and orange before normal fall color develops on unaffected plants. Leaf veins 

tend to be darker green than surrounding tissues (A: James A. Walla, Northern Tree Specialties, 

used with permission; B: Bruce Neill, Agriculture and Agri-food Canada, used with permission).

A B
56 USDA Forest Service RMRS-GTR-335. 2016.

Discolored leaves and stem dieback may also be associated with 

other declines caused by drought, 

cankers, bark beetles, and herbicides. At early onset, X-disease 

cannot be reliably differentiated from healthy plants or plants 

with other problems. With more 

advanced disease, the texture of 

leaves, the pattern of leaf discoloration, the combination of symptoms 

and signs, and the pattern of disease 

progression become more reliable, 

but confirmation requires serological or molecular diagnostics. 

Confirmation is generally justified 

only for research or to discern the 

role of other possible chokecherry 

problems.

Disease Cycle

Research in eastern and western 

United States indicates that the 

pathogen is usually transmitted 

to Prunus hosts by leafhoppers in 

mid- to late-summer when infected 

insects feed on the phloem in leaves 

of healthy plants. In larger plants, 

phytoplasmas are distributed 

systemically to all plant parts over 

a period of years. They survive 

over winter primarily in roots 

and are distributed to upper plant 

parts a few weeks after budbreak. 

Leafhoppers feed on phloem of 

infected leaves to acquire X-disease 

phytoplasmas, and can transmit 

the phytoplasmas after four to five 

weeks.

Damage

X-disease is the most limiting problem of chokecherry in plantings and native stands 

in the central and northern Great Plains. Growth and fruit production are reduced. Most 

infected plants gradually decline and have a shortened lifespan. In a study in eastern 

Nebraska, symptoms appeared on more than 80 percent of chokecherry plants within 

three years after X-disease was introduced to a plot, and mortality was more than 50 

percent within eight years. Damage to infected plants is less pronounced in the Prairie 

Provinces. In North Dakota, the most susceptible large shrubs die within 10 years of 

infection, and most infected plants die within 20 years of infection, compared to expected 

longevity of 70+ years. A small percentage of infected common chokecherry remains 

healthy for many years. Younger or more stressed plants are more quickly and seriously damaged than older or less stressed plants. Infected plants have reduced ability to 

recover from other stresses. Awareness of damage caused by X-disease limits the use of 

chokecherry in new plantings. Infected American plums are not damaged. Black cherry 

(Prunus serotina) and Japanese flowering cherry (Prunus serrulata) are apparently not 

susceptible. 

Figure 14-2—Increasing effect of X-disease in chokecherry. (A) In 1993, this severely diseased chokecherry 

plant had symptomatic red leaves when other plants still 

had green leaves. (B) The same plant was nearly dead 

by 1997 (James A. Walla, Northern Tree Specialties, 

used with permission).

B

A
USDA Forest Service RMRS-GTR-335. 2016. 57

Figure 14-3—Effects of X-disease. The disease can cause 

(A) fruit to remain red and pointed compared to the normal dark 

purple and round berries on unaffected plants, and (B) stunted, 

deformed flowers to be formed late in the season (Bruce Neill, 

Agriculture and Agri-food Canada, used with permission). 

Management

Promoting vigorous growth of infected plants (for example, with proper management 

of weeds, soil moisture, and soil fertility) will slow the rate of decline. Consider planting 

resistant or nonhost plants in areas where X-disease is a problem. American plum can 

be included in plantings containing susceptible hosts with confidence that the X-disease 

pathogen will not seriously damage it. However, the disease could be transmitted from 

American plum to susceptible plants. To reduce the spread of the disease by insect 

vectors, avoid establishing new plantings near any Prunus species. Removal of other 

susceptible plants such as wild chokecherries, from a zone (500 feet suggested for eastern 

United States) around desired susceptible plants, such as a stone fruit orchard, will reduce 

the rate of transmission. Clean cultivate or mow the area in and around plantings to 

reduce resting habitat for vectors. Mix one or more nonhost plants among Prunus species 

in new plantings.

Selected References

Peterson, G.W. 1984. Spread and damage of western X-disease of chokecherry in eastern 

Nebraska plantings. Plant Disease. 68: 103–104.

Walla, J.A.; Dai, W. 2006. X-disease of chokecherry. Tree Talk. [Edited online news 

magazine]. 2(5): 6–12. http://www.ag.ndsu.edu/trees/whatnew/Tree_Talk_Dec2006-2.

pdf. (February 27, 2015).

A

B
58 USDA Forest Service RMRS-GTR-335. 2016.

15. Biscogniauxia Canker of Oak

Judith O’Mara and Nicole Opbroek

Oaks (Quercus spp.) are an important forest and landscape tree species found throughout the Great Plains from the Dakotas to Texas. Biscogniauxia canker (pronounced 

‘Bisk-o-nee-ox-e-a’, formerly Hypoxylon canker) can cause substantial damage to oaks 

in wooded areas and urban landscapes. Disease severity is generally associated with 

stressful environmental and site conditions.

Hosts and Distribution

Biscogniauxia canker is caused by the fungi Biscogniauxia atropunctata

(syn. Hypoxylon atropunctatum) and B. mediterranea (syn. H. mediterraneum). In North 

America, both species of Biscogniauxia can be found as far north as Ontario, and as far 

south as Texas and Georgia. Biscogniauxia is also distributed across central and eastern 

states as well as in the western states of California, Oregon, and Washington. 

Outbreaks of Biscogniauxia canker are common in the Great Plains following multiyear periods of heat and drought, particularly in Kansas, Missouri, Oklahoma, and Texas. 

Biscogniauxia canker is primarily a problem on oaks in the red oak group, although it 

can also affect white oaks. Other hardwood species affected include maple (Acer spp.), 

hickory (Carya spp.), sycamore (Platanus occidentalis), elm (Ulmus spp.), hackberry 

(Celtis spp.), hornbeam (Carpinus caroliniana), and basswood (Tilia americana). 

Symptoms and Signs

Symptoms of Biscogniauxia canker resemble drought stress. Affected trees exhibit a 

general thinning of the canopy, yellowing and wilting of the foliage, and branch dieback. 

As the disease progresses, a mass of fungal tissue (stroma) is formed underneath the 

bark in the cambium. The pressure from this mass causes the bark to pop off, exposing a 

crusty, tan-colored mat of fungal tissue (fig. 15-1). As the outer layer of the stroma wears 

off, the canker turns silver (fig. 15-2) and then black and crusty, similar in appearance to 

asphalt (fig. 15-3). The black, crusty stroma is diagnostic for Biscogniauxia canker. 

Figure 15-1—Dusty-brown conidial stroma of 

Biscogniauxia canker on post oak (Q. stellata) 

(Jake Weber, Kansas State University, used 

with permission). 

Figure 15-2—Silver stroma of Biscogniauxia

canker on oak (Nicole Opbroek, Kansas Forest 

Service, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 59

Disease Cycle

Biscogniauxia canker can affect 

trees of any age. Sexual spores 

(ascospores) are spread by blowing 

wind, insects, or splashing rain. 

They enter trees through natural 

openings or wounds and the fungus 

can persist for many years within 

the sapwood and bark of healthy 

trees without causing disease symptoms. Vigorous trees can keep the 

fungus in check until a stress event 

triggers the onset of disease. Factors 

such as drought stress, fire, root 

injury, chemical damage, compacted 

soil or addition of fill soil, improper 

pruning, storm damage, or damage 

by insect pests may weaken the 

tree. These stress events in turn can 

reduce the water potential in the 

tree and predispose it to infection. 

Canker development often occurs in 

the year after a major stress event, 

although B. atropunctata can produce cankers within two months.

Biscogniauxia canker is favored 

by low moisture content within the 

tree and warm, dry growing conditions. Both Biscogniauxia species 

are adapted to dry conditions with 

optimal growth from 85 and to 

95 °F. Growth of B. atropunctata

can occur at temperatures up to 

104 °F. After infection, development of stromatic tissue causes 

bark to slough off, exposing a tan 

stroma containing asexual spores 

(conidia). As the stroma matures, 

the tan conidial layer wears off and 

exposes a silver stroma with a black 

ascospore-producing perithecial 

layer beneath. Old cankers appear 

black and crusty. Biscogniauxia 

mediterranea tends to produce 

small patches, whereas B. atropunctata produces cankers that range from a few inches to 

several yards along the length of the tree. As the fungus grows into the sapwood, it can 

coalesce into areas of yellowish decay with distinctive black zone lines (fig. 15-4) at the 

edge. As the disease progresses, infected trees can lose large strips of bark and die within 

one to two growing seasons. 

Damage

Historical reports indicate a link between periods of drought stress, oak decline, and 

the presence of Biscogniauxia canker. A drought during 1980 to 1981 in South Carolina 

was associated with lowered water tables and large areas of red oak mortality along the 

coastal highway. Almost all dead and declining oaks showed the distinctive tan-silver 

stromata associated with Biscogniauxia canker. Regional records suggest that a 1980 

Figure 15-3—Mature black Biscogniauxia stroma on 

oak (Nicole Opbroek, Kansas Forest Service, used with 

permission).

Figure 15-4—Black zone lines and wood decay caused 

by Biscogniauxia canker (Nicole Opbroek, Kansas 

Forest Service, used with permission).
60 USDA Forest Service RMRS-GTR-335. 2016.

drought in east central Arkansas resulted 

in heavy oak mortality, affecting 85 percent of the oak trees within a 494-acre 

area. Biscogniauxia canker was found 

on 95 percent of the dead oaks, as well 

as in oak trees exhibiting dieback. 

More recently, multiyear droughts 

have significantly affected oak populations in wooded areas and landscapes 

in eastern Kansas and Oklahoma and 

western Missouri. After a severe drought 

from 2010 to 2012, Biscogniauxia 

canker was observed by the author on 

10 percent of the oaks in two watersheds 

in northeast Kansas. A multi-county 

survey in central and eastern Oklahoma 

documented Biscogniauxia canker on 

a range of hardwood trees, including 

maple, goldenrain tree (Koelreuteria 

paniculata), hackberry, hickory, pecan (C. illinoinensis), American elm 

(U. americana), sycamore, and many 

species of oak. Maple trees (fig. 15-5) 

were severely affected by Biscogniauxia 

canker in Kansas, Oklahoma, Missouri, 

and Colorado during this same period. 

Management

There are no curative measures for trees with extensive Biscogniauxia canker. It 

might be possible to slow disease progression on trees by pruning affected limbs and 

improving the vigor of the tree with regular care. Remove severely affected trees down to 

ground level, as the disease can affect the remaining stump. Destroy firewood taken from 

infected trees so that the disease does not spread. Preventive strategies in timber stands 

include sanitation cutting of infected trees, removing infected debris from the area, and 

pruning trees with storm-damaged limbs. 

Selected References

Bassett, E.N.; Fenn, P. 1984. Latent colonization and pathogenicity of Hypoxylon 

atropunctatum on oaks. Plant Disease. 68: 317–319.

Chapela, H.I.; Boddy, L. 1988. Fungal colonization of attached beech branches. II. 

Spatial and temporal organization of communities arising from latent invaders in bark 

and functional sapwood, under different moisture regimes. New Phytologist. [Online]. 

110: 47–57. http://www.jstor.org/stable/2434789. (February 24, 2015). 

Farr, D.F.; Bills, G.F.; Chamuris, G.P.; Rossman, A.Y. 1989. Fungi on plants and plant 

products in the United States. St. Paul, MN: APS Press. 1252 p. 

Lewis, R., Jr. 1981. Hypoxylon spp., Ganoderma lucidum, and Agrilus bilineatus

in association with drought related oak mortality in the south. (Abstract). 

Phytopathology. 71: 890. 

Schoeneweiss, D.F. 1981. The role of environmental stress in diseases of woody plants. 

Plant Disease. 65: 308–314.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Figure 15-5—Biscogniauxia canker on maple 

(Jake Weber, Kansas State University, used with 

permission).
USDA Forest Service RMRS-GTR-335. 2016. 61

16. Botryodiplodia Diseases

Mark O. Harrell and Laurie J. Stepanek; 

revised from Jerry W. Riffle and Joseph M. Krupinsky, 

Botryodiplodia canker of elms; Glenn W. Peterson and 

Harrison L. Morton, Botryodiplodia disease of Russian-olive; and 

Robert Lewis, Jr., and Kenneth E. Conway, Botryodiplodia canker of 

sycamore (Riffle and Peterson 1986)

Trees weakened by drought, heat, or other stressful conditions in the Great Plains are 

often damaged by the fungi Botryodiplodia hypodermia and Lasiodiplodia theobromae 

(formerly B. theobromae), which cause cankers, dieback, and tree mortality. Siberian elm 

(Ulmus pumila), Russian-olive (Elaeagnus angustifolia), and sycamore (Platanus occidentalis) are commonly attacked. Additional information on L. theobromae can be found 

in chapter 20.

Hosts and Distribution

B. hypodermia (also known as B. ulmicola, Sphaeropsis hypodermia, and S. ulmicola) 

is found on American (U. americana), Siberian (U. pumila) and English (U. procera) 

elms in the United States. It occurs principally on Siberian elm in North Dakota, South 

Dakota, Nebraska, Kansas, Oklahoma, Minnesota, and Montana. 

L. theobromae (once known as Botryodiplodia theobromae, Diplodia gossypina, D. 

natalensis, D. theobromae, and B. rhodina) occurs around the world and attacks woody 

and herbaceous plants in at least 280 genera. In the Great Plains L. theobromae is most 

damaging to Russian-olive and sycamore, but will also attack ailanthus (Ailanthus altissima), American holly (Ilex opaca), flowering dogwood (Cornus florida), hickory (Carya

spp.), oak (Quercus spp.), persimmon (Diospyros virginiana), and walnut (Juglans spp.). 

In some tree species the fungus is only weakly pathogenic and may cause cankers only 

when the trees are severely stressed.

Symptoms and Signs

B. hypodermia cankers on Siberian elm develop a reddish-brown to black discoloration on the bark surface (fig. 16-1). Infected inner bark tissues turn reddish brown to 

brownish black (fig. 16-2) and become water soaked and soft. Sapwood beneath the 

Figure 16-1—Botryodiplodia hypodermia

canker on Siberian elm (James A. Walla, 

Northern Tree Specialties, used with 

permission).

Figure 16-2—Siberian elm stem with 

reddish-brown inner bark killed by 

Botryodiplodia hypodermia (James A. 

Walla, Northern Tree Specialties, used with 

permission). 
62 USDA Forest Service RMRS-GTR-335. 2016.

Figure 16-3—Loose bark caused by 

Botryodiplodia hypodermia on Siberian 

elm (James A. Walla, Northern Tree 

Specialties, used with permission).

Figure 16-4—Brown single-celled conidium of 

Botryodiplodia hypodermia (William R. Jacobi, 

Colorado State University, used with permission).

infected bark turns reddish brown to brownish 

black as well. The bark frequently splits longitudinally and may become loose (fig. 16-3). When 

the fungus girdles an infected branch or trunk, 

the foliage wilts and branch dieback or top kill 

occurs. Sprouts often develop below girdling 

cankers. Yellow leaves on infected American elm 

branches may somewhat resemble symptoms 

caused by Dutch elm disease.

L. theobromae in Russian-olive and sycamore 

typically kills bark and cambium in strips that 

may extend many feet along the trunk and 

branches. On Russian-olive the cankers may 

initially be inconspicuous and indicated only by 

the presence of small dead branches along the 

strip. Mortality of these small branches is most 

readily detected in the upper crown during July 

and August. Major branches and stems may have 

more than one strip of dead tissue, and the strips 

may spiral along the stems. Infected Russianolives may exude gum from stems and branches, 

but gum is not a reliable indicator of this disease 

because gumming also can be caused by other 

pathogens.

On sycamore, L. theobromae cankers may appear as tan to dark brown streaks in the 

normally green to white smooth bark. Cankers may develop cracks as they enlarge and 

age. Twig and branch dieback may occur either on an individual branch or throughout 

most of the crown. In cases where the trunk is affected, sprouts may grow from the base 

of the trunk (root collar) and on the trunk below the canker. Stressful environmental 

conditions, such as drought, can cause 

cankers to develop more rapidly and 

become more severe. 

Both B. hypodermia and L. theobromae fruit on killed bark. The fruiting 

bodies (pycnidia) appear as small black 

pimplelike structures embedded in the 

outer layer of dead bark. If pycnidia are 

cut through by shaving the bark, they 

often appear to have black walls and 

white contents. Pycnidia of both species 

produce asexual spores (conidia) that 

may be one- or two-celled, and clear 

or brown (fig. 16-4). B. hypodermia

conidia are 20 to 36 μm × 13 to 18 μm

in size. L. theobromae conidia are 20 to 

30 μm × 10 to 15 μm.

Disease Cycle

Conidia on dead bark are the primary inoculum for new infections. Most conidia are 

probably dispersed by splashing rain or wind, but may also be carried by insects and 

pruning tools. When conidia make contact with wounded stems, branch stubs, and other 

suitable infection courts, they germinate and colonize host tissues.

Colonization does not always result in a girdling canker. Susceptibility of the tree, 

virulence of the fungal strain, and environmental conditions are determining factors in 

canker formation. The most virulent strains can cause cankers in nonstressed trees, but 

the least virulent strains do not. Canker development is favored by high temperature and 

water stress.
USDA Forest Service RMRS-GTR-335. 2016. 63

Damage

The effects of infection can range from small, inconspicuous cankers to tree mortality. 

Small cankers have little effect on trees. Large cankers slow the growth rate and cause 

wood defects, which weaken stems and makes them more vulnerable to wind breakage. 

Dieback changes the form of individual trees and severe dieback may kill trees.

Management

Losses from Botryodiplodia cankers can be reduced by cultural practices. Avoid 

wounding stems and making branch stubs. Prune during late fall or winter when fungal 

colonization is reduced because of low temperatures. Prune dead, dying, or severely cankered branches from infected trees during winter or before spring rains to prevent fungal 

spores from splashing to new infection sites.

Because water stress favors infection and disease development, avoid planting trees 

on dry sites. Use a mulch bed of wood or bark chips around trees on dry sites to improve 

soil conditions. Trees in landscapes should be watered appropriately during dry periods. 

When possible, plant seedlings that are adapted to a particular geographic or climatic site. 

Select genetically improved stock with resistance to Botryodiplodia if available. 

Selected References

Ash, C.L. 1979. Pathogenicity of Botryodiplodia hypodermia on American elm, Ulmus 

americana., Fargo, ND: North Dakota State University. 146 p. M.S. thesis.

Filer, T.H., Jr. 1969. Sycamore cankers caused by Botryodiplodia theobromae. 

Phytopathology. 59: 76–78.

Krupinksy, J.M. 1981. Botryodiplodia hypodermia and Tubercularia ulmea in cankers on 

Siberian elm in northern Great Plains windbreaks. Plant Disease. 65: 677–678.

Lewis, R., Jr. 1978. Influences of infection court, host vigor, and culture filtrates on 

canker production by Botryodiplodia theobromae conidia in sycamore. Plant Disease 

Reporter. 62: 934–937.

Lewis, R., Jr.; Van Arsdel, E.P. 1978. Vulnerability of water-stressed sycamores to strains 

of Botryodiplodia theobromae. Plant Disease Reporter. 62: 62-63.

Peterson, G.W. 1976. Disease of Russian-olive caused by Botryodiplodia theobromae. 

Plant Disease Reporter. 60: 490–494.

Riffle, J.W. 1981. Cankers. In: Stipes, R.J.; Campana, R.J., eds. Compendium of elm 

diseases. St. Paul, MN: American Phytopathological Society: 34–42.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Thompson, G.E. 1951. Die-back of sycamore. Plant Disease Reporter. 35: 29–30.
64 USDA Forest Service RMRS-GTR-335. 2016.

17. Brown Rot of Stone Fruits

John Ball and James A. Walla; revised from David S. Wysong and 

James A. Walla (Riffle and Peterson 1986)

Common chokecherry (Prunus virginiana) and American plum (P. americana) are 

two common windbreak and native trees in the Great Plains. These trees are also used as 

ornamentals for their attractive spring flowers. The fruit from these trees is collected and 

used for jams and jellies. Other stone fruits commonly planted in the region include apricot (P. armeniaca), peach (P. persica), hybrid plum, and sour cherry (P. cerasus). These 

trees suffer from numerous pests, one of which is brown rot.

Hosts and Distribution

Brown rot on stone and pome fruits in North America is caused by several fungi such 

as Monilinia fructicola, M. amelanchieris, M. demissa, and M. laxa, with M. fructicola

likely predominating on the northern Great Plains. 

The disease affects primarily stone fruits, members of the genus Prunus. Chokecherry 

and American plum are the most commonly affected species on the northern Great Plains, 

whereas all stone fruits are affected in the warmer, more humid southern Great Plains. 

Less frequently, it can also be found affecting pome fruits, such as apple (Malus spp.), 

pear (Pyrus spp.), and serviceberry (Amelanchier spp.).

Symptoms and Signs

Brown rot has two main phases: blossom and shoot blight, affecting the flowers, 

leaves, and new shoots; and brown rot, affecting the fruit. The blossom and shoot blight 

phase begins in early spring. The brown rot phase involves the browning and decay of affected fruit, usually as the fruit ripens and after harvest, as a result of both shoot and fruit 

infection. A twig canker phase may also occur. 

Figure 17-1—Shoot of western sandcherry (Prunus pumila) 

infected with brown rot (Simeon Wright, Missouri Department 

of Conservation, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 65

Blossom blight begins with infection of anthers and pistils in the flowers, eventually 

spreading to kill the entire flower and the peduncle. Petals turn brown and water soaked, 

and then shrivel and wilt but remain hanging from the twig, often until midsummer. 

Shoot blight begins with infections of new leaves or shoots, with the infection usually 

spreading to girdle the shoot, causing the shoot to wilt and hang down (fig. 17-1). 

Immature fruit may be infected at any stage of development, but such infections are 

often latent. Infection of mature fruit is more common. Fruit decay usually begins from 

recent and latent infections, when sugar content increases as fruits near maturity. Decay 

first appears as tan-brown, almost circular spots. Within a few days, these spots coalesce 

and the entire fruit will turn a light brown with a gray powder on the surface (fig. 17-2). 

The hanging, discolored fruit will also have a semi-watery appearance before beginning 

to dry and harden. The dry, infected fruit may fall to the ground or shrivel and become 

mummified while still hanging from the twig. These mummified fruits generally drop 

later in the season although some may remain hanging for the winter. 

Twig cankers sometimes develop through expansion of blossom, shoot, or fruit infections. Infected twigs will have a sunken, oval or elliptical canker. As cankers girdle or 

nearly girdle twigs, the foliage distal to the canker will often become yellow and wilt, 

but remain attached. Gum will often ooze from the canker edge, most commonly with 

apricot. This gum “glues” infected flowers or fruit to the twig.

Signs of the pathogens are apothecia, in which sexual ascospores are produced, and 

sporodochia, on which asexual conidia are produced. The rarely seen apothecia are wineglass-shaped tan mushroomlike structures usually 10 to 15 mm across that arise in spring 

from fruit mummies on the ground. Sporodochia appear as ash-gray to tan tufts that may 

cover part or all of infected plant tissue under moist conditions at any time of the growing season and in storage. When abundant, sporodochia often form in concentric rings. 

Identification of many Monilinia species has been difficult, but gene sequences have been 

described and proven useful.

Disease Cycle

Windblown ascospores from apothecia and splashed, windblown, or insect-carried 

conidia from mummified fruit or twig cankers are the inoculum sources for infection of 

opening flowers and expanding shoots in spring. Some Monilinia species (including M. 

fructicola) produce repeated secondary cycles of conidia throughout the season; others 

produce only one secondary cycle of conidia. Secondary conidia infect immature and 

mature fruit, primarily through wounds, but also through stomata and hair sockets. Fruits 

Figure 17-2—Formation of ash-gray mold on surface of fruit infected with brown 

rot (Simeon Wright, Missouri Department of Conservation, used with permission). 
66 USDA Forest Service RMRS-GTR-335. 2016.

infected by both groups will develop brown rot, mostly as they ripen, and the fruit will 

become mummified and overwinter to continue the disease cycle. Fungus survival is 

reduced in colder climates for fruit mummies that hang on tree branches, but not for those 

on the ground.

Damage

The disease is more common in warmer, and wet, climates, such as those of the eastern United States. Brown rot can result in the loss of more than half the fruit in an orchard 

if not treated, particularly during summers with above-normal precipitation and humid 

conditions. The disease often is more noticeable as blossom and shoot blight on chokecherry. Both phases, blossom and shoot blight and brown rot, appear on American plum.

Management

Sanitation is a key practice in reducing this disease. Mummified fruit on and under the 

tree as well as infected twigs should be removed and destroyed before spring. Remove 

wild chokecherry or plum trees from the vicinity. Also, improve drying conditions in and 

around susceptible trees, for example by pruning stone fruits for an open center; increasing the distance between susceptible trees; tilling or mowing ground cover; and pruning, 

thinning, or removing overhanging and surrounding trees. Control vectoring insects that 

make or visit wounds in the fruit, such as fruit flies (Drosophila spp.), oriental fruit moth 

(Grapholita molesta), and plum curculio (Conotrachelus nenuphar). Fungicide treatments are often necessary.

Brown rot is usually managed with fungicides during two time periods: the spring to 

reduce blossom and shoot blight and the summer to reduce brown rot on fruit. One to 

three fungicide sprays are applied to manage blossom blight: the first as the flowers open, 

the second at full bloom, and the third at petal fall. A similar schedule is used for shoot 

blight, beginning as the leaves unfurl. Spring may be the only season that treatment is 

needed if the ornamental value of the flowers and foliage is the only concern. If the fruit 

is important, however, then summer fungicide applications are made beginning about 

three weeks before harvest. Several applications may be needed during fruit ripening, 

and fungicide labels should be checked for preharvest intervals. If brown rot infestation remains too high, control of latent infections may be needed in future years. Many 

populations of brown rot fungi have developed resistance to individual fungicides, so use 

different fungicide groups in alternating applications for disease control.

Infection can easily spread from infected to healthy fruit during storage. To reduce 

storage losses, harvest before fruit is overripe, avoid wounding the fruit, sort out any 

distorted and discolored fruit, cool the fruit promptly after harvest, and periodically sort 

stored fruit to remove those with decay.

Selected References

Holb, I.J. 2004. The brown rot fungi of fruit crops (Monilinia spp): III. Important features 

of their disease control. International Journal of Horticultural Science. 10: 31–48.

Holb, I.J. 2008. Brown rot blossom blight of pome and stone fruits: symptom, disease 

cycle, host resistance, and biological control. International Journal of Horticultural 

Science. 14: 15–21.

Luo, Y.; Michailides, T.J. 2001. Factors affecting latent infection of prune fruit by 

Monilinia fructicola. Phytopathology. 91: 864–872.

Northover, J.; Biggs, A.R. 1995. Effects of conidal concentration of Monilinia 

fructicola on brown rot development in detached cherries. Canadian Journal of Plant 

Pathology—Revue Canadienne de Phytopathologie. 16: 30–36

Xu, X.M.; Robinson, J.D. 2000. Epidemiology of brown rot (Monilina fructigena) on 

apple: infection of fruits by conidia. Plant Pathology. 49: 201–206.
USDA Forest Service RMRS-GTR-335. 2016. 67

18. Black Knot of Cherry and Plum

Mark O. Harrell and James T. Blodgett; revised from David S. 

Wysong and Mark O. Harrell (Riffle and Peterson 1986)

Black knot, caused by the fungus Apiosporina morbosa, can significantly reduce fruit 

production and kill small branches in heavily infected trees.

Hosts and Distribution

Black knot occurs throughout the United States. It affects cherries, plums, and most 

other members of the genus Prunus. Common hosts in the Great Plains include bird 

(Prunus padus), bitter (P. emarginata), black (P. serotina), choke (P. virginiana), mahaleb 

(P. mahaleb), Nanking (P. tomentosa), pin (P. pensylvanica), sand (P. pumila), sour 

(P. cerasus), and sweet (P. avium) cherries; apricot (P. armeniaca) and peach (P. persica); 

and American (P. americana), Canada (P. nigra), and domestic plums (Prunus spp.).

Symptoms and Signs

The fungus produces elongate woody swellings (galls) or knots on twigs, branches, 

and small stems (figs. 18-1 and 18-2). The knots are usually about one to eight inches 

long and up to one inch thick, but some may reach one foot or more in length. On large 

branches and stems, infections can result in cankerous galls or rough bark, or both. 

Scattered black masses of fungal tissue (stromata) that produce fruiting bodies also may 

be present. Knots become greenish and soft in late spring about one year after infection, 

but become black and hard over time (fig. 18-2). Old knots may be covered with a white 

to pink mycoparasitic fungus (Trichothecium roseum) (fig. 18-3), and insects often tunnel 

into the knots.

Figure 18-1—Black knot on chokecherry (James A. Walla, 

Northern Tree Specialties, used with permission).
68 USDA Forest Service RMRS-GTR-335. 2016.

When mature, knot surfaces are covered with hard black fungal fruiting bodies (pseudothecia) composed entirely of stromata. The inner knot is composed of both host and 

fungal tissues. Pseudothecia produce two-celled club-shaped, olivaceous spores (ascospores) measuring 13 to 18 μm × 4.5 to 7.5 μm.

Disease Cycle

Ascospores (sexual spores) of the fungus are discharged in spring from fruiting bodies 

on the surface of the knots. Sporulation may occur from the time that tips of green leaves 

emerge from buds until shoot growth stops. Sporulation is often greatest from the pink/

white blossom stage to two weeks after bloom. Rain is required for spore discharge, and 

spores are carried by rainsplash and wind. Germinating spores penetrate unwounded 

surface tissue of current-season growth and may also infect older tissues through wounds. 

Infection is most severe when prolonged moist conditions are accompanied by temperatures between 55 and 77 °F.

Knots appear several months after infection. Depending on host species, cultivar 

susceptibility, and length of the growing season, some knots are visible by late summer; 

others do not appear until the following spring. Knots that appear during fall or spring 

continue growing through the next summer. Conidiospores (asexual reproductive spores) 

are produced on developing knots beginning about when ascospore discharge ends from 

mature knots and continuing into midsummer. These spores are considered to have little 

role in infection. 

At least one year and usually two years are required before new knots produce mature 

fruiting bodies. Fruiting bodies on the old knot die after releasing ascospores in spring, 

but the fungus often grows into adjacent wood the year before sporulation and produces 

ascospores there the next year. Some infections may expand for several years, potentially 

expanding into large branches and stems.

Figure 18-3—Old black knot swelling partially 

covered by a white parasitic fungus (James T. 

Blodgett, U.S. Forest Service).

Figure 18-2— Old (black) and new (olive green) 

black knot swellings (Aaron D. Bergdahl, North 

Dakota Forest Service, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 69

Damage

Severely infected trees can have little value due to physical and aesthetic effects. 

Heavy black knot infections can significantly reduce plant vigor and fruit production. 

Older branches beyond knots frequently die. New branches beyond knots often do not 

develop or new shoots may leaf out and wilt in early summer.

Management

The disease is kept in check by less than optimal infection conditions; thus, management is usually not needed. Unbalanced host-pathogen phenology, partial host resistance, 

and the presence of mycoparasites can also reduce the need for extensive disease 

management.

Damage from black knot can be reduced by pruning. Damage can also be mitigated by 

removing or pruning infected wild hosts in nearby wooded areas and fence rows within 

600 ft of the desired host trees. Make pruning cuts three to four inches or more below 

the knot, as the fungus may extend beyond the swelling. The best times to prune are late 

winter and early spring, because knots are most easily seen before the tree has leaves and 

because the source of spores should be removed before the next infection season begins. 

Pruned knots on the ground are another source of spores that can cause new infections if 

left in the area, so gather and dispose of all pruned knots before April 1.

Because the disease usually has a two-year life cycle, stems should be closely inspected and knots should be removed for two consecutive years. Infections from the previous 

year will often be small or not visible when mature knots are pruned the first year. In 

subsequent years, less removal is required to prevent disease buildup.

If black knot is severe within 200 yards of a planting site, do not plant Prunus species, 

or select resistant species and cultivars when possible. The pathogen is often host specialized, and a species or cultivar thought to be resistant may be susceptible to a pathogen 

form present in the area that is different from where the plant was tested, or the plant may 

become infected if new pathogen forms move into the area.

Fungicide sprays can provide some protection, but the protection is not sufficient 

if sanitation and pruning are not included. Fungicides registered in some states for use 

against black knot include chlorothalonil, copper hydroxide, mancozeb, and thiophanatemethyl. Full protection would require the fungicide to be present throughout the 

sporulation period. However, if infection pressure is not severe, then protection from the 

pink/white blossom stage until two weeks after bloom may be sufficient. Follow label 

instructions for application timing, precautions, and other information.

Selected References

Northover, J.; McFadden-Smith, W. 1995. Control and epidemiology of Apiosporina 

morbosa of plum and sour cherry. Canadian Journal of Plant Pathology. 17: 57–68.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs, 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Snover, K.L.; Arneson, P.A. 2002. Black knot. The Plant Health Instructor. [Online]. 

doi:10.1094/PHI-I-2002-0926-01. http://www.apsnet.org/edcenter/intropp/lessons/

fungi/ascomycetes/pages/blackknot.aspx. (April 24, 2014).

Zhang, J.X.; Fernando, W.G.D.; Remphrey, W.R. 2005. Genetic diversity and structure of 

the Apiosporina morbosa populations on Prunus spp. Phytopathology. 95: 859–866.
70 USDA Forest Service RMRS-GTR-335. 2016.

19. Black-Spot Nectria Canker

Marcus B. Jackson; revised from Tubercularia Canker of Siberian 

Elm and Russian-olive, by Joseph M. Krupinsky and James A. Walla 

(Riffle and Peterson 1986)

Black-spot Nectria canker causes dieback and mortality of trees weakened by drought, 

freeze injury, root damage, poor site adaptability, and other stresses. The disease is caused 

by the fungus Nectria nigrescens and is usually identified by mature fruiting bodies of its 

anamorph (asexual stage) Tubercularia ulmea, which resemble raised black spots on the 

bark of affected trees and shrubs.

Hosts and Distribution

N. nigrescens is found on more than 30 species of maple (Acer); birch (Betula); 

peashrub (Caragana); bittersweet (Celastrus); Russian-olive (Elaeagnus); ash 

(Fraxinus); honeylocust (Gleditisia); creeper (Parthenocissus); ninebark (Physocarpus); 

poplar, aspen, and cottonwood (Populus); plum, peach, nectarine, and cherry (Prunus); 

buckthorn (Rhamnus); sumac (Rhus); currant (Ribes); willow (Salix); elderberry 

(Sambucus); buffaloberry (Shepherdia); mountain-ash (Sorbus); lilac (Syringa); linden 

(Tilia); and elm (Ulmus). This fungus occurs in Europe and temperate North America 

with extensive damage occasionally observed in northern Great Plains states and the 

Prairie Provinces. The most severe damage in the Great Plains has been in nonnative 

Siberian elm (U. pumila) and Russian-olive (E. angustifolia), as well as in honeylocust 

(G. triacanthos) trees planted outside their native range.

Symptoms and Signs

Flags (recently killed branches with 

dead leaves still attached) may be the first 

indication that black-spot Nectria canker is 

affecting a tree (fig.19-1). Closer examination near the point of infection may show 

oval to elongate areas of sunken bark, which 

are sometimes darkened on small, thinbarked branches (fig. 19-2). Gum exudates 

may occur on some hosts, such as Russianolive. Removing bark within the sunken 

area will show dead, discolored (often dark 

brown to black) cambium and outer sapwood 

indicative of a branch or stem canker (fig. 

19-3). Many different pathogens can cause 

cankers on the dozens of N. nigrescens hosts, 

so identification of the pathogen by examination of fruiting structures is required. 

N. nigrescens-caused canker is usually 

confirmed by identifying the anamorph, 

which fruits more readily both in nature 

and under culture than the sexual stage (teleomorph). Cushion-shaped fruiting bodies 

(sporodochia) of T. ulmea are fairly common 

on N. nigrescens-killed branches (fig. 19-4), 

but are not always present. They are 0.25 to 1.7 mm high and 0.3 to 1.7 mm wide with a 

stalk that may not be readily evident as it is often covered by the bark surface. The tops 

can be white, cream, tan, orange, red, brown, or black at first, but will typically turn black 

within two to three weeks after sporodochia form. Stalks are white to whitish red, or, 

rarely, dark red. One-celled, clear (hyaline), straight, or slightly curved ellipsoidal to cylindrical conidia measure 4.6 to 8.4 μm × 1.5 to 3.0 μm (fig. 19-5). They form on curved 

(sometimes straight or coiled) hyaline conidiophores on top of sporodochia (fig 19-6). 

Figure 19-1—Flagging of Siberian elm caused 

by black-spot Nectria canker (Marcus B. 

Jackson, U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 71

Figure 19-3—Canker margin exposed by peeling bark on 

a Siberian elm (Marcus B. Jackson, U.S. Forest Service). 

Figure 19-2—Sunken bark in a canker 

on the stem of a two-year-old Siberian 

elm inoculated with Nectria nigrescens (Marcus B. Jackson, U.S. Forest 

Service).

Figure 19-4—Fruiting bodies (sporodochia) of Tubercularia 

ulmea (anamorph of Nectria nigrescens); lower right, 

cross-section of a sporodochium (Marcus B. Jackson, U.S. 

Forest Service).

Figure 19-5—Spores of Tubercularia 

ulmea (anamorph of Nectria nigrescens) 

(Marcus B. Jackson, U.S. Forest Service).

T. ulmea fruiting bodies can be 

confused with those of the coralspot Nectria canker anamorph, 

but the former blacken with age, 

whereas T. vulgaris typically 

retains a lighter color. 

Teleomorphic fruiting bodies 

of N. nigrescens are infrequently 

found in nature. They are red to 

reddish-brown single or clustered 

perithecia on raised stromata and 

are sometimes clustered around 

the base of a sporodochium. 

Perithecia are 0.27 to 0.42 mm 

high and 0.24 to 0.41 mm wide 

with aseptate to three-septate 

(mostly one-septate) ascospores that measure 

10.5 to 22.0 μm × 2.5 to 8.0 μm (figs. 19-7 

and 19-8). They are easily confused with N. 

cinnabarina fruiting bodies but have minor microscopic differences. Rapid field discrimination 

between the two Nectria species is by the color 

of mature sporodochia. When fruiting bodies 

are not present, the fungus can be cultured from 

wood chips taken from the edge of a young 

canker. T. ulmea can produce sporodochia after 

two to six weeks on potato dextrose agar at 

room temperature.
72 USDA Forest Service RMRS-GTR-335. 2016.

Disease Cycle

N. nigrescens overwinters as mycelium and fruiting bodies in cankered bark. Although windborne 

spores of the teleomorph are important for pathogen 

adaptability and long-distance spread, they are probably insignificant for local disease outbreaks owing 

to their rare occurrence. Most infections are caused 

by asexual spores (conidia) that are held in place by 

a viscous material until liberated by water. They are 

spread by rainsplash, by adhering to birds or insects, 

or on pruning tools. Conidia infect bark wounds 

caused by hail, wind, freeze damage, snow breakage, 

animals, cultivation, pruning, and possibly herbicides. 

The fungus invades the sapwood, cambium, and 

living portions of the bark. It is considered a weak 

pathogen, causing more damage to stressed trees. In 

otherwise healthy trees, only small cankers usually 

develop, which will seal over with woundwood tissue 

in the next growing season. N. nigrescens generally 

grows more quickly in weakened branches and stems, 

resulting in more girdling infections that cause flagging and provide a large amount of substrate for more 

conidial production.

Figure 19-6—Spore-producing structures (sporophores) of Tubercularia ulmea (anamorph of 

Nectria nigrescens) (Marcus B. Jackson, U.S. 

Forest Service). 

Figure 19-7—Nectria nigrescens fruiting bodies 

(perithecia) (Marcus B. Jackson, U.S. Forest 

Service).

Figure 19-8—Nectria nigrescens 

ascospores within a spore sac 

(ascus) (Marcus B. Jackson, U.S. 

Forest Service). 
USDA Forest Service RMRS-GTR-335. 2016. 73

Damage

Although N. nigrescens has a large host range, widespread damage is rare and is largely restricted to stressed trees. Root damage from transplanting or excavating near trees, 

drought stress, freeze injury, and chronic herbicide exposure have all been implicated in 

significant damage caused by N. nigrescens. Extensive damage sometimes occurs on trees 

planted beyond their native range, such as Russian-olive, Siberian elm, and honeylocust 

in the Great Plains. Trees that are well adapted to their environment generally have little 

or no damage from N. nigrescens. Therefore, ongoing problems with this pathogen may 

indicate trees are poorly adapted to a given site or a site requires some form of mitigation 

to reduce the stress on affected species. 

Management

Impacts of black-spot Nectria canker can be reduced with proper woody plant selection, plant stress mitigation, and sanitation. Select well-adapted species suitable to soils, 

weather conditions, and maintenance capabilities of the site. Some cultivars have been 

developed with improved tolerance to cold, drought, and common diseases and insects 

that weaken trees and shrubs. These cultivars may be less prone to N. nigrescens damage. 

Planting adapted species and cultivars may reduce damage from this disease. Because 

healthy trees and shrubs are less susceptible to damage, maintain proper irrigation and 

fertilization and avoid injury from soil compaction, root disturbance, and herbicides. 

Manage other diseases and insects when feasible. 

Avoid unnecessary bark wounds. Complete pruning during cool, dry periods when 

spore dispersal and germination are limited, making infection less likely. Dead branches 

and trees can be burned or removed from the site to reduce inoculum for future infections. See chapters 20 and 21 for more specific information about managing N. nigrescens

on host species.

Selected References

Hirooka, Y.; Rossman, A.Y.; Samuels, G.J.; Lechat, C.; Chaverri, P. 2012. A monograph 

of Allantonectria, Nectria, and Pleonectria (Nectriaceae, Hypocreales, Ascomycota) 

and their pycnidial, sporodochial and synnematous anamorphs. Studies in Mycology. 

71(1): 1–210.

Jackson, M.B. 1997. Influence of host and chemical treatments on cankers caused by 

Tubercularia ulmea. Fargo, ND: North Dakota State University. 91 p. M.S. thesis.

Jackson, M.B.; Stack, R.W. 2002. Effects of dicamba herbicide on Tubercularia ulmea

canker development. Journal of Arboriculture. 28(2): 94–98.

Krupinsky, J.M. 1981. Botryodiplodia hypodermia and Tubercularia ulmea cankers on 

Siberian elm in northern Great Plains windbreaks. Plant Disease. 65(8): 677–678.

Sinclair, W. A.; Lyon, H. H. 2005. Tubercularia canker and dieback. In: Diseases of trees 

and shrubs, 2nd ed. Ithaca, NY: Cornell University Press: 178. 
74 USDA Forest Service RMRS-GTR-335. 2016.

20. Canker and Dieback of Russian-Olive 

Gerard C. Adams, Ned Tisserat, and William R. Jacobi

Russian-olive (Elaeagnus angustifolia) is used in windbreaks, shelterbelts, and 

landscape plantings. It has been planted widely in windbreaks in the northern Great 

Plains because it survives rigorous environmental conditions. It is particularly tolerant of drought and salinity. In some regions it has become an invasive plant in riparian 

environments with major negative impacts on native plants. In the Great Plains the tree is 

susceptible to several fungal canker diseases: 

•  Phomopsis canker caused by Phomopsis arnoldiae (syn. P. elaeagni)

•  Botryodiplodia canker caused by Lasiodiplodia theobromae (formerly Botryodiplodia 

theobromae) 

•  Black-spot Nectria canker caused by Nectria nigrescens (asexual state Tubercularia 

ulmea) and also called Tubercularia canker. 

Hosts and Distribution

Phomopsis canker is the most important canker disease of Russian-olive throughout 

the Great Plains, the North Central states, and the Northeast. The disease was first found 

in Missouri in 1963 and subsequently in Illinois, Ohio, Delaware, Michigan, and New York. 

The only other recorded host of P. arnoldiae is black walnut (Juglans nigra) nursery seedling 

stock in Indiana. Other species of Phomopsis cause cankers and dieback on many plant 

species. Many other plants also harbor opportunistic Phomopsis species that are common endophytes. Inoculation tests can aid in distinguishing aggressive pathogens from opportunists. 

Botryodiplodia canker is common on many woody plants. Lasiodiplodia theobromae 

is an opportunist latent pathogen, and is an endophyte with worldwide distribution. The 

pathogen commonly causes cankers on trees suffering from environmental stress such as 

drought. More information on Botryodiplodia canker can be found in chapter 16.

Black-spot Nectria canker on Siberian elm (Ulmus pumila), Russian-olive, and other 

common hosts is discussed in chapters 19 and 21. N. nigrescens (a member of the N. cinnabarina species complex) is a common pathogen of trees suffering from environmental 

stress such as freezing before full dormancy has been reached. 

Symptoms and Signs

Recently killed branches with dead leaves still attached (flags) indicate the presence of 

a canker that has girdled the stem or branch. Phomopsis and black-spot Nectria cankers 

are elongate, girdling, reddish-brown to purplish-black, and mostly on small branches, 

while those of Botryodiplodia cankers are long, narrow strips mostly on stems. There is 

an obvious canker margin in the bark tissue, especially in younger tissues, visible on the 

bark surface for Phomopsis and Tubercularia cankers (fig. 20-1), but only by excising 

the bark for Botryodiplodia cankers. The sapwood immediately beneath the bark canker 

is brown, and this browning may extend beyond the margin of the canker. Frequently 

gummosis occurs around the canker margins. Phomopsis and black-spot Nectria cankers 

often develop on current-year shoots. Site of development of Botryodiplodia cankers on 

Russian-olive is not known, but most cankers are found on larger stems. Young shoots 

are girdled quickly, wilting the new silvery foliage. Cankers also have been found on 

branches up to four inches in diameter, and larger for Botryodiplodia cankers. Black-spot 

Nectria and Phomopsis cankers often look similar, and all three cankers may be present in 

the same area or tree, particularly in the central Great Plains.

The asexual fruiting bodies (pycnidia) of Phomopsis form on infected bark in the same 

season the bark is killed. Pycnidia with multiple locules are numerous and clustered in 

cankers, erupt through the bark, and measure approximately 500 μm × 275 μm. Asexual 

spores (conidia) in pycnidia, or exuding in moist weather, are of two types known as 

alpha and beta. Alpha conidia are short (5 to 11μm long), ovoid, blunt, and straight 

(fig. 20-2); beta conidia are long (15 to 26 μm), filiform, and curved (fig. 20-3). Both 

spore types are produced on plant tissue and sometimes in culture. The beta conidia act as 

spermatia and are not infective. 
USDA Forest Service RMRS-GTR-335. 2016. 75

Figure 20-1—Discolored bark of Phomopsis canker on young Russian-olive 

branch with asexual fruiting bodies (pycnidia) beginning to erupt through bark in 

cankered area (Ned Tisserat, Colorado State University, used with permission).

Figure 20-2—Alpha conidia of Phomopsis spp. 

are oval and colorless, 5-11 μm long (Paul Bachi, 

University of Kentucky, Bugwood.org, used with 

permission).

Figure 20-3—Beta conidia of Phomopsis spp. 

are filiform and curved, 15-26 μm long (Bruce 

Watt, University of Maine, Bugwood.org, used 

with permission).

The pycnidia of L. theobromae also can form on infected bark in the same season the 

bark is killed. Additionally, they are often observed on twigs and branches that died a few 

years earlier. The pycnidia erupt through bark separately (fig. 20-4) or as groups in fused 

clusters. The fused clusters may superficially appear as one fruiting body with multiple 

locules, but individual pycnidia have one locule. The compound fruiting body is large 

(1-5 mm), whereas the individual pycnidia are about 250 μm in diameter. The conidia 

begin as colorless single cells and then become brown two-celled bodies ornamented with 

striations (figs. 20-5 and 20-6). Lasiodiplodia is readily distinguished from Phomopsis by 

the large ellipsoid conidia that measure 20 to 30 μm × 10 to 15 μm (fig. 20-6). 

The asexual fruiting bodies (sporodochia) of black-spot Nectria canker are usually 

observed on twigs and branches that have died the previous year but may appear in the 

same year as infection, particularly in branches that are rapidly killed. Sporodochia are 

distinctive cushions erupting through the bark. Initially they may be cream, tan, reddish, 

brown, or black but turn black about two or more weeks after eruption (fig. 20-7). In 

North Dakota and South Dakota, N. nigrescens is more frequently isolated from the cankers than Phomopsis or Lasiodiplodia. The conidia are colorless and oval (fig. 20-8).

When fruiting bodies are not present, the pathogens are confirmed by culturing 

wood chips taken from the edge of the canker, on potato dextrose agar or similar media. 

Phomopsis produces white fluffy aerial hyphae and pycnidia in stromata. Lasiodiplodia 
76 USDA Forest Service RMRS-GTR-335. 2016.

Figure 20-8—The conidia of the black-spot 

Nectria canker pathogen are produced on 

the sporodochia and are colorless and oval 

(Bruce Watt, University of Maine, Bugwood.

org, used with permission).

Figure 20-6—The mature conidia of L. 

theobromae are two-celled, dark brown, 

and large (20-30 × 10-15 μm) (Photo: Cesar 

Calderon, USDA APHIS PPQ, Bugwood.org, 

used with permission).

Figure 20-7—The black-spot fruiting bodies (sporodochia) are usually visible on the bark of the infected 

branch (Bruce Watt, University of Maine, Bugwood.

org, used with permission).

produces gray aerial hyphae and pycnidia in 

stromata. Nectria produces sporodochia usually within two months of incubation on potato 

dextrose agar at room temperature. 

Disease Cycle

The life cycle of these parasites has not 

been demonstrated, but studies of Phomopsis,

Botryodiplodia, and black-spot Nectria cankers 

in other hosts indicate that conidia infect 

through wounds and become established as endophytes in the sapwood. While the fungi are 

in the previous-year sapwood and sometimes 

the leaves, new growth remains free of the fungi for a period and is susceptible to infection. It 

is assumed that new growth becomes infected 

by conidia. It is not known whether infections 

can occur throughout the growing season. 

Cankers develop within a few days following natural infection or artificial inoculation. 

Figure 20-4—The asexual fruiting bodies 

(pycnidia) of the Botryodiplodia canker pathogens erupt through the bark separately or in 

clusters (Paul Bachi, University of Kentucky, 

bugwood.org, used with permission)..

Figure 20-5—A cross-sectional cut through a 

pycnidium demonstrates that the large conidia 

(20-30 × 10-15 μm) of L. theobromae are colorless and one-celled at first, before becoming 

dark brown and two-celled (Cesar Calderon, 

USDA APHIS PPQ, Bugwood.org, used with 

permission). 
USDA Forest Service RMRS-GTR-335. 2016. 77

Sporulation on infected tissue also follows quickly on black-spot Nectria and Phomopsis 

cankers, but is several weeks delayed for Botryodiplodia canker. Black-spot Nectria and 

Phomopsis cankers are annual and delimited by an active host response. Cankers on older 

tissues occur during periods of stress when the host defense response is compromised. 

Infection on older tissue is believed to result from expansion of endophytic colonies of 

the previously latent pathogen. Botryodiplodia cankers are typically perennial with no 

apparent host response.

Damage

In the northern Great Plains, these canker diseases contribute to the decline of 

Russian-olive health, and damage to the function of shelterbelts and windbreaks that include the species. The canker diseases are usually restricted to trees under environmental 

stress and the stress may be present in subsequent years. Such trees exhibit increasing 

dieback of branches, resulting in decreasing health and aesthetic appearance of the 

canopy over the years. Both disease incidence and severity increase with age until the 

quality of plant appearance is no longer acceptable and the plants become increasingly 

susceptible to other pathogens and pests. 

Infected Russian-olive and black walnut nursery stock may suffer severe damage 

from Phomopsis canker, especially in the North Central states. Terminal dieback of black 

walnut nursery stock and outplanted saplings leads to multiple-stemmed trees. Careful 

inspection of healthy-appearing Russian-olive ornamental plants may reveal that as much 

as 52 percent have the disease, as was found in a study in southeastern Michigan. It is 

recommended that diseased nursery stock be disposed of by being buried, burned, or 

composted away from production plants. 

Management

Because healthy trees are less susceptible to infection and damage, manage trees for 

optimum vigor. If possible, water trees during prolonged droughts. Establish new plantings on good sites with vigorous planting stock and control weeds. Reduce chances of 

infection by preventing wounds to the bark. Because infected or dead branches of trees 

are a source of fungal inoculum, remove the branches from the site and burn them. Prune 

infected branches back to the nearest living branch six inches beyond the canker. Prune 

trees during dry weather, and disinfect pruning tools after each cut. 

Selected References

Jackson, M.B. 1997. Influence of host and chemical treatments on cankers caused by 

Tubercularia ulmea, Fargo, ND: North Dakota State University. 91 p. M.S. thesis.

Krupinsky, J.M. 1986. Cankers on Russian olive in windbreaks in North Dakota and 

South Dakota. Plant Disease. 70: 981.

Krupinsky, J.M.; Frank, A.L. 1986. Effect of water stress on Tubercularia canker 

of Russian olive. Pp. 171-172. In Proc. International Symposium on Windbreak 

Technology, D.L. Hintz and J.R. Brandle, eds. June 23-27, Lincoln, NE. Great Plains 

Agricultural Council Publication No. 117. 269 p.

Maffei, H.M.; Morton, H.L. 1983. Phomopsis canker of Russian-olive in southeastern 

Michigan. Plant Disease. 67: 964–965.

Morehart, A.L.; Carroll, R.B.; Stuart, M. 1980. Phomopsis canker and dieback of 

Elaeagnus angustifolia. Plant Disease. 64: 66–69.

Peterson, G.W. 1967. Botryodiplodia disease of Russian-Olive. Phytopathology. 57(8): 

825–826.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Stannard, M.; Ogle, D.; Holzworth, L.; Scianna, J.; Sunleaf, E. 2002. History, biology, 

ecology, suppression, and revegetation of Russian-olive sites (Elaeagnus angustifolia, 

L.). Technical Notes. Plant Materials No. 47. Bozeman, MT: U.S. Department of 

Agriculture, Natural Resources Conservation Service. 14 p.
78 USDA Forest Service RMRS-GTR-335. 2016.

21. Cankers and Root Collar Rot of Honeylocusts

William R. Jacobi and James A. Walla 

Branch and stem cankers are among the most serious problems of honeylocusts and 

can be caused by various fungal pathogens. Thyronectria canker is caused by Pleonectria 

austroamericana, formerly Thyronectria. Black-spot Nectria canker, formerly called 

Tubercularia canker, is caused by Nectria nigrescens, the recently described sexual 

stage of Tubercularia ulmea. More information on this canker can be found in chapter 

19. Coral-spot Nectria canker is caused by Nectria cinnabarina, the sexual stage of 

Tubercularia vulgaris. Root collar rot of honeylocust acts like a girdling canker at the soil 

line; the pathogen is not known.

Hosts and Distribution

Thyronectria canker occurs on native thorny honeylocust (Gleditsia triacanthos 

var. triacanthos), all cultivars of thornless and podless honeylocust (G. triacanthos var.

inermis) and oriental honeylocust (G. japonica). The fungus P. austroamericana also 

colonizes and produces spores on dead tissue of several other hardwoods including 

American elm (Ulmus americana), white ash (Fraxinus americana), mockernut hickory 

(Carya tomentosa), willow (Salix spp.), and bur oak (Quercus macrocarpa). The disease 

has been reported from Colorado and Oklahoma eastward to Massachusetts and Georgia. 

The disease is common in the central United States, where it can cause major damage. It 

is uncommon in the Northeast and in Great Lake states such as Minnesota and Wisconsin, 

and it has not been found in the northernmost states and provinces of the Great Plains, so 

fungus distribution may be restricted by colder climates.

Black-spot Nectria canker is common on many hardwood and coniferous trees 

and shrubs. The fungus N. nigrescens is found on honeylocusts, maples (Acer spp.), 

Russian-olive (Elaeagnus angustifolia), winged euonymus (Euonymus alatus), green ash 

(F. pennsylvanica), American (U. americana) and Siberian elm (U. pumila), and many 

other woody plants. Among honeylocusts, those planted outside of their native range are 

the most susceptible to damage from this disease. The disease is found commonly from 

Colorado eastward and from the northern Great Plains states northward to Manitoba and 

Alberta.

Coral-spot Nectria canker is found on many tree species, and honeylocusts are a 

common host of N. cinnabarina in some areas of the Great Plains. The disease can be 

found from the northern to the southern Great Plains but is the least common canker 

disease on honeylocusts of these three described.

Root collar rot is a common disease on honeylocusts in the drier regions of the 

Great Plains, where irrigation practices can keep the soil saturated at the base of the tree. 

Occurrence of the disease is also related to weather conditions that apparently promote 

the disease in the dormant season. Thus, the disease can be seen girdling trees for several 

years in a row, and then there can be several years without return of the disease.

Symptoms and Signs

Symptoms of the four diseases include dieback of affected branches, reduced foliage, 

yellow foliage, premature fall coloration, and early leaf drop.

•  Cankers range from slightly flattened surfaces to distinctly sunken areas with large 

ridges of woundwood at the canker margin. Loose bark and woundwood at the tree 

base can indicate root collar rot. Thin-barked areas of stems and branches may have a 

red-yellow discoloration associated with the canker. Usually there is no discoloration 

where the bark is thicker. Wood beneath infected bark turns wine-red to yellow for 

Thyronectria canker.

•  Honeylocust root collar rot symptoms include discolored and loose bark and discolored wood at the soil line. Early fall coloration of a portion of the tree may indicate 

a large amount of damage. Small drops of gum on the stem or root sprouting usually 

indicate girdling. Loose bark and discolored (yellow to brown instead of white) wood 

just below the bark indicate initial collar rot and are the most indicative symptoms. 
USDA Forest Service RMRS-GTR-335. 2016. 79

Signs of the pathogen often allow for better disease identification than do symptoms.

•  Thyronectria cankers have irregularly shaped clumps of stroma (fig. 21-1, 21-2, 

21-3) containing irregularly shaped chambers lined with conidiophores (fig. 21-4) 

(stromatic conidiomata) instead of neatly organized round pycnidia with necks. 

Stromatic conidiomata are light pink or yellow-brown when fresh and become 

blackened with age. Perithecia are yellowish brown or reddish gray with dark, often 

black, shining apical regions and also darken with age (fig. 21-5). Both fruiting body 

types are usually found in natural openings such as lenticels in thick-bark areas and 

scattered on bark surfaces in thin-bark areas. Asexual fruiting bodies (pycnidia) 

exude milky masses of conidia (fig. 21-6). Conidia are hyaline, ovoid to ellipsoid, 

Figure 21-1—Thyronectria canker on a honeylocust with asexual fruiting bodies (pycnidia) growing 

through bark lenticels (William R. Jacobi, Colorado 

State University, used with permission).

Figure 21-2—Asexual fruiting bodies (pycnidia) 

in Thyronectria canker in cracked bark (William 

R. Jacobi, Colorado State University, used with 

permission). 

Figure 21-3—Asexual fruiting bodies (pycnidia) in Thyronectria canker (William R. 

Jacobi, Colorado State University, used with 

permission). 

Figure 21-6—Creamy-white asexual spores (conidia) oozing from fruiting bodies of Thyronectria 

canker (William Jacobi, Colorado State 

University, used with permission).

Figure 21-4—Pleonectria 

austroamericana conidia 

and conidiogenous cells 

(Yuuri Hirooka, University 

of Maryland, used with 

permission).

Figure 21-5—Perithecia in 

Thyronectria canker (Yuuri Hirooka, 

University of Maryland, used with 

permission).
80 USDA Forest Service RMRS-GTR-335. 2016.

Figure 21-08—Pleonectria austroamericana sexual spores 

(ascospores), mostly in asci (William Jacobi, Colorado State 

University, used with permission).

one-celled, and 1.6 to 4.2 μm × 0.8 to 2.7 μm in size. Sexual fruiting bodies (perithecia) develop in rounded clusters in a stroma, and are smooth-surfaced and egg-shaped 

with a short neck. Spore sacs (asci) contain eight ellipsoid to pear-shaped ascospores 

that are hyaline to pale yellow with both transverse and longitudinal septa and range 

in size from 8 to 16 μm × 5 to 9 μm (fig. 21-7, 21-8). Cultures of P. austroamericana

on potato dextrose agar have somewhat waxy margins with orange, slimy centers 

containing masses of conidia (fig. 21-9).

•  Black-spot Nectria cankers (fig. 21-10, 21-11) appear very similar to Thyronectria 

cankers, and are best distinguished by cultural characteristics, morphology of the 

asexual fruiting structures, and ascospore morphology. The asexual sporodochia are 

creamy to peach colored when fresh, but normally turn dark brown to black within 

a few days of drying conditions (fig. 21-12). They usually form exposed on the bark 

surface or erupt from a stroma under a thin layer of bark. A fresh squash mount of the 

sporodochia will show curved conidiogenous cells (fig. 21-13, 21-14). Perithecia are 

reddish brown and form singly or in groups of up to 20 on a raised fungal stroma and 

can be found clustered at the base of sporodochia (fig. 21-15, 21-16). Ascospores are 

ellipsoid to fusiform, smooth walled, one- to three-septate, and 13.5 to 18.0 μm × 3.5 

to 5.5 μm in size (fig. 21-17). N. nigrescens cultures are white and fluffy.

Figure 21-10—Black-spot 

Nectria canker with black sporodochia on honeylocust (William 

R. Jacobi, Colorado State 

University, used with permission). 

Figure 21-11—Black-spot 

Nectria canker at branch crotch 

(William R. Jacobi, Colorado 

State University, used with 

permission).

Figure 21-7—Pleonectria 

austroamericana asci with 

multicelled sexual spores 

(Yuuri Hirooka, University 

of Maryland, used with 

permission).

Figure 21-9—Culture 

plate of Pleonectria austroamericana with orange, 

slimy center (Yuuri Hirooka, 

University of Maryland, used 

with permission).
USDA Forest Service RMRS-GTR-335. 2016. 81

Figure 21-12—Black-spot Nectria canker with old darkened sporodochia on 

bark (William R. Jacobi, Colorado State 

University, used with permission). 

Figure 21-13—Sporodochia with curved 

conidiogenous cells (spore-bearing structures) and conidia of Nectria nigrescens 

(William Jacobi, Colorado State University, 

used with permission).

Figure 21-16—Creamy colored sporodochia surrounded 

by red fruiting bodies (perithecia) in black-spot Nectria 

canker (Yuuri Hirooka, University of Maryland, used with 

permission).

Figure 21-15—Clumps of fruiting bodies 

(perithecia) in black-spot Nectria canker (William R. Jacobi, Colorado State 

University, used with permission).

Figure 21-14—Curved conidiogenous 

cells and spores (conidia) of Nectria 

nigrescens (Yuuri Hirooka, University 

of Maryland, used with permission).

Figure 21-17—Sexual spores 

(ascospores) of Nectria nigrescens (Yuuri Hirooka, University of 

Maryland, used with permission).

•  Coral-spot Nectria cankers (fig. 21-18) appear similar to the two other cankers, 

but N. cinnabarina may form sporodochia that are creamy to bright coral colored 

when young (usually for several weeks) and tan, brown or black after overwintering 

(fig. 21-19). Perithecia form in late summer singly or in groups of up to 15 and are 

bright red to reddish brown (fig. 21-20, 21-21). Ascospores are one- to two-septate, 

fusiform, and 14.0 to 17.5 μm × 4.0 to 5.5 μm (fig. 21-22), similar to N. nigrescens

but unlike those of P. austroamericana. N. cinnabarina cultures are white and fluffy.

Disease Cycle

The three canker-causing fungi overwinter as mycelium and fruiting structures on 

infected trees. Conidia presumably are spread by rain and ascospores by wind. Infections 

take place through wounds and natural openings. Because these fungi can exist as saprophytes, they can become established on dead wood such as branch stubs or wound edges. 
82 USDA Forest Service RMRS-GTR-335. 2016.

Figure 21-18—Discolored honeylocust bark 

from coral-spot Nectria canker (William R. 

Jacobi, Colorado State University, used with 

permission). 

Figure 21-19—Sporodochia of coral-spot 

Nectria canker in honeylocust (William R. 

Jacobi, Colorado State University, used with 

permission). 

Figure 21-20—Red fruiting bodies (perithecia) of coral-spot Nectria canker (William 

Jacobi, Colorado State University, used with 

permission).

Figure 21-21—Red fruiting bodies (perithecia) 

surrounding white sporodochia of coral-spot 

Nectria canker (Yuuri Hirooka, University of 

Maryland, used with permission).

As pathogens, the fungi grow in the cambium and outer xylem, where they eventually kill 

the cambium and surrounding cells. Trees, or affected parts, die because of cambial death 

and possibly vascular dysfunction. Sporodochia can form within one month after tissue 

is colonized, and are abundant on bark of dying or dead trees. Perithecia form in affected 

areas but are not as common or abundant as pycnidia or sporodochia. 

The disease cycle of honeylocust root collar rot involves interactions with pathogenic 

soil organisms through unknown processes. Frequent irrigation that keeps the soil wet 

at the tree base allows soil microorganisms to kill the bark and cambium just below soil 

line. P. austroamericana, N. cinnabarina, or N. nigrescens then may infect the weakened 

tree above the area previously killed by root collar rot and cause large cankers that can be 

mistaken for the primary disease.

Damage

Cankers at the tree base are usually fatal. Cankers at the main stem or branch crotch 

may cause complete girdling, depending on the tree’s health. Stressed trees cannot defend 

against the fungi, whereas vigorous trees should be able to stop the fungus invasion and 

recover. The amount of damage by the canker diseases across the United States is not 

known, but they are found on many hosts and in particular on honeylocust in urban, rural, 

and windbreak plantings. Root collar rot can kill trees rapidly.

Management

Honeylocust cankers and root collar rot should be prevented rather than controlled. 

Physical damage and wounds should be avoided or properly cleaned and allowed to dry. 

Susceptibility to these pathogens is increased by a variety of stresses, including drought, 

overwatering, herbicide damage, and restricted area or restricted oxygen for root growth. 

Trees should be watered adequately but not excessively. Frequent irrigation of turf is a 
USDA Forest Service RMRS-GTR-335. 2016. 83

Figure 21-22—Sexual spores (ascospores, left) of Nectria cinnabarina (Yuuri Hirooka, 

University of Maryland, used with permission). Asexual spores (conidia, right) and conidiogenous cells of Nectria cinnabarina (Yuuri Hirooka, University of Maryland, used with 

permission).

factor that creates conditions favoring root collar rot. This disease can be prevented by 

keeping the soil at the base of trees as dry as possible. Place sprinklers and sprinkler 

heads far away from trees to minimize the amount of water falling on the tree stem and at 

the tree base. Remove flowers, turf, or other vegetation that requires frequent irrigation 

from around the tree’s base and replace with small gravel or organic mulch. When planting honeylocust trees, place the top of the root ball six inches above grade to allow for 

drainage away from the tree trunk. Black-spot Nectria canker rarely occurs on ‘Northern 

Acclaim’ honeylocust in the northern Great Plains, likely owing to its superior cold hardiness. ‘Sunburst’ honeylocust is the most susceptible to Thyronectria canker. ‘Imperial,’ 

‘Skyline,’ and thornless selections are more resistant.

Pruning infected branches can reduce the chances of other infections. Trees should 

be pruned in cool, dry weather when the presence of fungal spores is reduced and when 

exposed bark will be sealed over the soonest. Small cankers may be scribed out if the tree 

is reasonably vigorous.

Selected References

Crowe, F.; Starkey, D.; Lengkeek, V. 1982. Honeylocust canker in Kansas caused by 

Thyronectria austro-americana. Plant Disease. 66: 155–158.

Hirooka, Y.; Rossman, A. Y.; Samuels, G.J.; Lechat, C.; Chaverii, P. 2013. A monograph 

of Allantonectria, Nectria and Pleonectria (Nectriaceae, Hypocreales, Ascomycota) 

and their pycnidial, sporodochial and synnematous anamorphs. Studies in Mycology. 

71: 1–210.

Hudler, G.W.; Oshima, N. 1976. The occurrence and distribution of Thyronectria austroamericana on honeylocust in Colorado. Plant Disease Reporter. 60: 920–922.

Jacobi, W.R. 1989. Resistance of honeylocust cultivars to Thyronectria austro-americana. 

Plant Disease. 73: 805–807.

Jacobi, W.R. 1992. Seasonal effects on wound susceptibility and canker expansion 

on honeylocusts inoculated with Thyronectria austro-americana. Journal of 

Arboriculture. 18: 288–293.

Jacobi, W.R.; Riffle. J.W. 1989. Effects of water-stress on Thyronectria canker of 

honeylocusts. Phytopathology. 79: 1333–1337.

Riffle, J.W.; Peterson, G.W. 1986. Thyronectria canker of honeylocust: influence of 

temperature and wound age on disease development. Phytopathology. 76: 313–316.
84 USDA Forest Service RMRS-GTR-335. 2016.

22. Crown Gall

Simeon Wright; revised from Michael W. Ferguson and 

Richard Dorset (Riffle and Peterson 1986)

Crown gall is a tumor-forming plant disease caused by Rhizobium radiobacter (formerly Agrobacterium tumefaciens) and related bacterial species occurring around the 

world. It is a serious issue for the nursery industry and affects both broadleaf and conifer 

species (fig. 22-1). Crown gall can sometimes be challenging to diagnose because of 

similarities with other galls and the prevalence of nonpathogenic strains of Rhizobium. 

Hosts and Distribution

More than 600 host plants in more than 90 

families are susceptible to crown gall, but natural 

symptom development has not been observed in 

the field for all species. Species and strains of 

Rhizobium are highly species specific. Apple and 

crabapple (Malus spp.), cottonwood (Populus

spp.), plum and cherry (Prunus spp.), and willow 

(Salix spp.) are frequently reported as hosts in the 

Great Plains, although other species are reported 

as well. 

Symptoms and Signs

Individual swellings (galls) develop on roots 

or stems, often at the base (root collar) of the tree 

(fig. 22-2). Several galls may be present along 

a stem or root (fig. 22-3). Gall diameter varies 

from less than 1 inch to more than 12 inches 

(fig. 22-4). The exterior surface of galls is usually 

rounded with a rough, irregular surface that darkens with age (fig. 22-5). The interior can 

be spongy or hard and solid with irregular structure. Galls caused by insects, mites, and 

fungi may look similar but often contain holes or chambers or have a nodular exterior.

Disease Cycle

R. radiobacter is a soil-inhabiting bacterium infecting fresh wounds that result from 

mechanical or environmental injury such as propagation, pruning, freezing, insect or 

nematode damage. After entering the plant, bacterial DNA is transferred to the host, 

stimulating increased cellular growth and division within that region of the host tissue 

through the production of phytohormones. The swollen tissue that develops forms the 

gall. Trees may be stunted when the woody tissue is disrupted, preventing flow of water 

Figure 22-2—Crown gall symptoms at the root 

collar (Aaron D. Bergdahl, North Dakota State 

University, used with permission).

Figure 22-3—Multiple galls along tree 

roots (William R. Jacobi, Colorado State 

University, used with permission).

Figure 22-1—Crown gall on juniper 

seedling (William R. Jacobi, Colorado 

State University, used with permission). 
USDA Forest Service RMRS-GTR-335. 2016. 85

or nutrients. Chewing insects or tools can carry the 

bacteria from plant to plant. 

Long-distance movement is 

also possible by transport of 

infected nursery stock or by 

contaminated soil, irrigation 

water, or tools such as pruning or grafting equipment. 

Symptoms may not appear 

for several weeks after infection, depending on weather 

conditions. Gall development 

is favored by temperatures 

near 72 °F. Bacteria return 

to the soil from galls and 

can persist for long periods 

of time, colonizing root 

surfaces of nonsusceptible 

plants or as a latent infection 

on susceptible hosts. Crown 

gall is favored by soils of 

neutral or higher pH.

Damage

Infection of large trees is 

infrequent and may not result 

in economic loss. Infection 

of nursery stock can be quite 

extensive, especially when 

inoculation takes place during propagation. Small trees 

may become stunted and 

more susceptible to environmental stress. In most states there is no tolerance for nursery 

stock infected with crown gall; all infected plants are destroyed. 

Management

Always use disease-free planting stock and propagation materials. Extension guides 

list resistant species that can be planted in infested areas. Avoid mechanical wounds on 

young trees or bushes. Sterilize cutting instruments frequently by using a disinfectant 

such as 70 percent alcohol. Remove and destroy all infected plants and rotate with a nonsusceptible crop (corn, small grains, other grass) for two to four years. As a preventive 

pre-planting treatment, nursery stock can be dipped in a suspension containing a related 

antagonistic bacterial species R. rhizogenes (syn. Agrobacterium radiobacter) (Strain 

K84 or K1026) to protect against damage from sensitive strains of R. radiobacter. 

Selected References

De Cleene, M.; De Ley, J. 1976. The host range of crown gall. The Botanical Review. 42: 

389–466.

Kado, C.I. 2010. Plant bacteriology. St. Paul, MN: American Phytopathological Society 

Press. 336 p.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Vicedo, B.; Peñalver, R.; Asins, M.J.; López, M.M. 1993. Biological control of 

Agrobacterium tumefaciens, colonization, and pAgK84 transfer with Agrobacterium 

radiobacter K84 and the tra-

 mutant strain K1026. Applied Environmental 

Microbiology. 59(1): 309–315.

Figure 22-4—Large aerial gall on cottonwood (William R. Jacobi, 

Colorado State University, used with permission). 

Figure 22-5—Rough gall exterior at cottonwood tree base 

(William R. Jacobi, Colorado State University, used with 

permission).
86 USDA Forest Service RMRS-GTR-335. 2016.

23. Cryptodiaporthe (Dothichiza) Canker of Populus

Species 

Michael E. Ostry and Jared M. LeBoldus; revised from David W. 

Johnson and Robert W. Stack (Riffle and Peterson 1986)

Disfiguring and often lethal stem cankers caused by Cryptodiaporthe populea affect 

primarily black poplars (Populus nigra varieties) in nurseries, plantations, and landscape 

plantings in North America.

Hosts and Distribution

Cryptodiaporthe canker (also known as Dothichiza canker) caused by the fungus 

C. populea is a widespread disease affecting poplars in nurseries, plantations, and 

landscape plantings throughout the Northern Hemisphere and in South America. 

Susceptibility to the disease varies by poplar species, but hybrids in sections Aigeiros 

(cottonwoods, black poplars) and Tacamahaca (balsam poplars) are the most frequently 

and severely affected. Lombardy poplar (P. nigra var. italica), which was commonly used 

for ornamental line plantings, is so highly susceptible that its life is greatly shortened and 

it is no longer recommended for use. The disease is not significant on aspen (P. tremuloides) in native stands.

Symptoms and Signs

Young, developing cankers, often 

at branch axils, begin as discolored 

patches of bark on twigs, branches, 

and stems of trees of any age. Cankers 

enlarge and the bark within the canker 

margins becomes slightly sunken. 

Raised, dark, pimplelike fruiting 

structures (pycnidia) containing spores 

(conidia) of the asexual state of the 

fungus (Discosporium populeum; syn. 

Dothichiza populea and Chondroplea 

populea) develop within the infected 

bark (fig. 23-1). Later, the bark cracks 

and extensive woundwood often 

develops around the perennial cankers, resulting 

in rough, warty areas on larger branches and stems 

(fig. 23-2). Adventitious shoots often develop below cankers that have girdled stems. 

Disease Cycle

Conidia, lemon shaped and 9 to 11 μm × 7 to 8 

μm in size (fig. 23-3), are released in cream-colored 

tendrils from pycnidia within the canker margins 

during wet weather throughout the growing season 

(fig. 23-4). The conidia are dispersed by rainsplash 

or inadvertently carried by insects or birds to 

infection sites such as buds, leaf scars, lenticels, 

and various wounds on adjacent trees. Ascospores, 

produced by the sexual state of the fungus, develop 

in small, flask-shaped fruiting bodies (perithecia), 

and are not as commonly encountered as the conidia. These spores presumably are also infective 

Figure 23-1—Young branch Cryptodiaporthe canker 

with fruiting bodies (pycnidia) protruding through 

the discolored bark (Michael E. Ostry, U.S. Forest 

Service).

Figure 23-2—Perennial, rough stem 

Cryptodiaporthe cankers with extensive woundwood (Michael E. Ostry, 

U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 87

and involved in spread of the fungus. The 

pathogen overwinters as spores and mycelium 

within cankers. After infection, cankers can 

rapidly expand in the bark of trees under 

moisture stress, but their growth may be arrested by tree defensive mechanisms, such 

as woundwood production, with the return 

of normal growing conditions. However, 

because the fungus can persist in the bark 

and wood, the cankers may begin to expand 

when trees are again exposed to stressful 

conditions. 

Damage

The fungus causes bark necrosis, cankers, 

dieback, and death of affected trees in nurseries and established tree plantings. Affected 

trees are disfigured and the longevity of 

highly susceptible trees used in landscape 

plantings is greatly reduced. Cankers girdle or 

weaken tree branches and stems, and invasion 

by decay fungi into the open cankers results 

in reduction in wood quality and stem breakage (fig. 23-5). The disease is most severe on 

trees under moisture stress and trees stressed 

or damaged by other abiotic and biotic 

agents. 

Management

Poplar varieties known to be vulnerable to Cryptodiaporthe 

canker should not be planted. In addition, avoid poplar cultivars and clones highly susceptible to leaf diseases, because 

premature defoliation stresses trees. The resulting leaf scars 

provide infection sites for C. populea during the growing 

season. Only vigorous, disease-free stock should be planted. 

The fungus can be harbored on hardwood cuttings and nursery 

stock without visible symptoms. Apply nursery practices that 

maintain tree vigor such as prudent fertilization and weed control. Remove diseased trees and plant material to reduce fungal 

inoculum. Store and handle planting stock to avoid moisture 

stress and conditions that favor the development of the 

pathogen. Avoid wounding trees and prevent moisture stress 

to minimize the risk of infection and damage by the fungus in 

landscape plantings and plantations. 

Selected References

Ostry, M.E.; McNabb, H.S., Jr. 1982. How to identify and prevent injury to poplars 

caused by Cytospora, Phomopsis, and Dothichiza. HT-57. St. Paul, MN: U.S. 

Department of Agriculture, Forest Service, North Central Forest Experiment Station. 

6 p.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Figure 23-5—Stem breakage at Cryptodiaporthe 

canker (Michael E. Ostry, 

U.S. Forest Service).

Figure 23-3—Lemon-shaped asexual spores 

(conidia) of the Cryptodiaporthe canker. Scale 

bar is 10 μm (Michael E. Ostry, U.S. Forest 

Service).

Figure 23-4—Cream-colored tendrils of 

asexual spores (conidia) exuded from fruiting bodies (pycnidia) in a Cryptodiaporthe 

canker (Michael E. Ostry, U.S. Forest 

Service).
88 USDA Forest Service RMRS-GTR-335. 2016.

24. Cryptosphaeria Canker of Cottonwood and Aspen

James J. Worrall, Kelly S. Burns, and Jennifer Juzwik; revised from 

Thomas E. Hinds and Jerry W. Riffle (Riffle and Peterson 1986)

Cryptosphaeria canker, caused by the fungus Cryptosphaeria ligniota (syn. C. populina), is a destructive disease of aspen and other poplars (Populus spp.). Although the 

fungus had been collected from Colorado as early as 1897, it was not associated with a 

canker disease until 1969. The anamorph (asexual stage) is a Cytosporina species (often 

referred to as Libertella species). 

Hosts and Distribution

Quaking aspen (P. tremuloides) is the principal host of C. ligniota, but the disease also 

occurs on other Populus species of the Great Plains, including narrowleaf (P. angustifolia), eastern (P. deltoides), and plains (P. deltoides ssp. monilifera) cottonwoods; 

Lombardy (P. nigra var. italica), balsam (P. balsamifera), and black (P. nigra) poplars; 

and some hybrid poplars. The disease is widely distributed and has been found throughout much of North America and in Europe.

Symptoms and Signs

Cryptosphaeria cankers are usually much longer and narrower than other cankers of 

aspen and may spiral snakelike around the stem, following the grain of the underlying 

wood (fig. 24-1). For this reason, the disease is sometimes referred to as “snake canker.” 

Cankers may be only two to four inches wide but 10 ft long. They may extend for 

much of the stem length, but more often occur primarily in the portion of the stem with 

live branches. Bark recently killed near the margin of the canker becomes discolored 

light brown to orange. Reddish-brown fluid may bleed from the margin. Within one to 

two years, dead inner bark becomes stringy, black, and sooty, and contains diagnostic 

small (0.5 to 2.0 mm), lenticular, light-colored spots (fig. 24-2). Annual woundwood 

formation may be visible but does not restrict canker growth. Dead bark adheres tightly 

to cankers.

The disease is a canker rot, 

which means that the fungus 

advances in and decays wood, 

causing cankers where the infection reaches the cambium and 

bark. Wood is stained in hues of 

gray, brown, yellow, orange, and 

even pink. The decay is brownish and mottled and is probably 

a type of soft rot similar to white 

rot, as is typical of wood-decaying ascomycetes. Wood staining 

can extend longitudinally three 

feet or more above and below 

the canker margin. The pathogen 

can be isolated readily from 

cankers and underlying wood. 

Sexual fruiting bodies (perithecia) form in narrow patches 

up to 0.5 inch wide and 1 foot 

long on the surface of bark that 

has been dead at least one year 

Figure 24-1—Cryptosphaeria canker killing an aspen. The 

canker spirals around the stem up to the live crown (James 

J. Worrall, U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 89

(fig. 24-3). The fungus readily fruits on branches and smooth bark, but perithecia are 

uncommon both on small trees and on thick, corky bark of old trees. Light-orange fruiting bodies (acervuli) of the Cytosporina (Libertella) stage are occasionally found near the 

canker margin on aspen. 

Cytospora species frequently infect and fruit at the margin of Cryptosphaeria cankers 

and quickly colonize trees after they die, so the cause of death may be erroneously attributed to Cytospora canker. For this reason, and because the pathogen may not fruit 

reliably, diagnosis should be made by looking for the lenticular, light-colored areas in 

bark and the staining in the sapwood. 

Figure 24-2—Light-colored lenticular spots that frequently appear in older dead 

bark of Cryptosphaeria cankers. Live bark is visible at the left-most margin of the cut 

(James J. Worrall, U.S. Forest Service).

Figure 24-3—Sexual fruiting bodies (perithecia) of Cryptosphaeria ligniota (numerous small, dark dots) (James J. Worrall, U.S. Forest Service).
90 USDA Forest Service RMRS-GTR-335. 2016.

Disease Cycle

Infection occurs primarily through branch and trunk wounds. Initially, the pathogen 

colonizes the heartwood and sapwood (xylem), causing discoloration and decay, but the 

fungus eventually grows out from the wood to the cambium and bark (phloem), killing 

the cambium and causing a canker. Wood staining can extend longitudinally three ft or 

more above and below the canker margin. Trees usually die before the canker completely 

girdles the stem.

Black perithecia may appear scattered on bark that has been dead for at least one year. 

The perithecia develop singly in a grayish pseudostroma made of dense fungal hyphae 

and dead bark just beneath the surface. Sexual spores (ascospores) are forcibly ejected 

from perithecia during wet weather and are presumably dispersed by wind. Spores from 

the Cytosporina stage likely function as fertilizing elements rather than causing infection. 

Damage

Cryptosphaeria canker is an important killer of aspen. In one Colorado survey, 

83 percent of 30 surveyed sites had Cryptosphaeria canker, and 26 percent of aspen 

mortality was attributed to the disease. It seemed to kill mostly small to midsize trees. 

Cryptosphaeria was the canker most frequently found on trees in diameter classes ranging 

from three to six inches and was found only on trees less than 11 inches in diameter. It 

is estimated that eight percent of aspen in Colorado have decay caused by C. ligniota. 

Cryptosphaeria canker is one of a number of wound-infecting cankers that lead to 

mortality following partial cutting of aspen stands and also in developed recreation sites 

established in aspen. 

Mortality usually occurs in small-diameter trees after only a few years and before the 

canker girdles the stem. A large volume of sapwood can be killed, causing more damage 

than the canker itself. On larger trees, infections may start in branches, and then spread 

into the stem, where they can grow up and down and girdle other branches, killing the 

tree one branch at a time. The importance of this disease on Populus species of the Great 

Plains is largely unknown. 

Management

Wounding should be prevented. However, many infections result from natural infection courts, and practical means of preventing such infections are not known. Pruning 

diseased, dying, and dead branches on high-value trees may reduce impacts by preventing 

the fungus from spreading into the stem. Development of recreation sites in aspen stands 

is strongly discouraged because aspen is prone to many diseases that can lead to stand 

deterioration. 

Selected References

Hinds, T.E. 1976. Aspen mortality in Rocky Mountain campgrounds. Res. Pap. RM-164. 

Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain 

Forest and Range Experiment Station. 20 p.

Hinds, T.E. 1981. Cryptosphaeria canker and Libertella decay of aspen. Phytopathology. 

71(11): 1137–1145.

Juzwik, J.; Nishijima, W.T.; Hinds, T.E. 1978. Survey of aspen cankers in Colorado. Plant 

Disease Reporter. 62(10): 906–910.

Sinclair, W.A.; Lyon, H.H. 2005. Cryptosphaeria canker of aspen. In: Diseases of trees 

and shrubs. 2nd ed. Ithaca, NY: Cornell University Press. 660 p.

Walters, J.W.; Hinds, T.E.; Johnson, D.W.; Beatty, J.S. 1982. Effects of partial cutting 

on diseases, mortality and regeneration of Rocky Mountain aspen stands. Res. Pap. 

RM-240. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky 

Mountain Forest and Range Experiment Station. 12 p.
USDA Forest Service RMRS-GTR-335. 2016. 91

25. Cytospora Canker of Hardwoods

Gerard C. Adams and William R. Jacobi 

Cytospora canker, caused by various species of Cytospora (sexual forms, Valsa and 

Leucostoma), is a common disease of many hardwoods including poplars (Populus spp.), 

willows (Salix spp.), maples (Acer spp.) elms (Ulmus spp.), alders (Alnus spp.), ash 

(Fraxinus spp.), peach (Prunus persica), mountain-ash (Sorbus spp.), quince (Cydonia

spp.), chokecherry (Prunus virginiana), serviceberry (Amelanchier arborea), and various 

woody plants in the Great Plains. Cytospora species are generally latent pathogens that 

form cankers on hosts subjected to environmental stresses. 

Hosts and Distribution

Susceptible hosts include those growing on poor sites or injured by drought, frost and 

winter injury, sunscald, fire, herbicides, mechanical damage, severe pruning, insects, 

or disease such as fire blight (see chapter 26). In alders, cankering is associated with 

unusually hot weather. The latent pathogens often colonize senescing branches and twigs 

without forming evident cankers. For example, Cytospora species colonize nearly every 

senescing branch and twig on maples planted in suitable urban sites. In unsuitable urban 

sites the same Cytospora species will cause damaging, often lethal, perennial cankers 

on the living trunks of maples. Occasionally, Cytospora species can be opportunistic 

invaders of bark and twigs that have been killed by other causes, and these species are 

generally not listed in host indexes. Differentiating between a latent pathogenic and a 

saprophytic Cytospora species is generally possible by inoculating hosts during early or 

late dormancy or when the plant is slightly stressed such as by drought, or by inoculating 

excised living twigs. Summer inoculations during active host defense response often fail 

to form cankers.

Cytospora canker is widespread in North America and other regions. Many Cytospora

species have been inadvertently distributed worldwide with their preferred host as endophytes in propagation cuttings. Resistance to Cytospora canker varies among hosts, but in 

general there are no commercially available resistant genotypes.

Symptoms and Signs

Cytospora species cause branch dieback and cankers on trees of any age. Cankers 

on trunks and limbs are often elongate, slightly sunken, discolored areas in the bark 

(fig. 25-1). Bark often splits along the canker margin because of woundwood formation 

by the host. The fungus may quickly girdle and kill twigs without forming cankers. 

Symptoms vary with host species and stage of disease development. Bark above infected 

cambium may appear sunken and yellow, brown, reddish brown, gray, or black. Diseased 

inner-bark and cambium turn reddish brown to black, and become watery and odorous as 

the tissues deteriorate. Gummosis occurs in Prunus species. Wood below the cambium is 

stained brown. 

Fruiting bodies develop in dead bark. Asexual 

fruiting bodies (pycnidia) 

form first and appear 

as small pustules on 

smooth bark (figs. 25-1, 

25-2). Pycnidia are less 

conspicuous on rough 

bark. Superficial cuts in 

cankered bark may expose small (generally 0.5 

to 1.5 mm diameter) dark 

pycnidia. Pycnidia generally have white deposits 

of dried spores encircling 

Figure 25-1—Cytospora canker on stem of peach showing shrunken 

discolored bark (Gerard C. Adams, University of Nebraska, used with 

permission).
92 USDA Forest Service RMRS-GTR-335. 2016.

Figure 25-2—Cytospora canker on stem of poplar showing shrunken bark and minute pustules 

of fruiting bodies (William R. Jacobi, Colorado 

State University, used with permission).

Figure 25-3—Asexual (pycnidia; small gray 

structures) and sexual (perithecia; large black 

structure) fruiting bodies of Cytospora on aspen 

bark (William R. Jacobi, Colorado State University, 

used with permission).

the openings (ostioles). During moist weather, yellow 

to reddish-brown spore masses or tendrils may exude 

from pycnidia (fig. 25-4). Dried tendrils may be the 

first sign noticed on some cankered tissues. The spores 

(conidia) are clear (hyaline), sausage-shaped (allantoid), 

one-celled, and generally 3 to 5 μm × 1.0 to 1.5 μm. 

Sexual fruiting bodies (perithecia) are sometimes found 

in bark after pycnidia mature in the year after death of 

the cambium. On the host, the fungal stroma forms larger 

pustules (2 to 8 mm in diameter), each containing perithecia (each about 0.3 to 0.5 mm in diameter) in groups 

of six to 20 with necks converging toward the surface of 

a disk of whitish to dark brownish tissue. The disk (upper 

surface of the stroma) breaks through the bark (fig. 25-3). 

Sometimes the necks extend through the disk as visible 

beaks. Ascospores from perithecia are hyaline, allantoid, 

one-celled, and about 7 to 12 μm × 1.5 to 2.5 μm.

Disease Cycle

Sexual and asexual spores of Cytospora species are 

infective, especially when freshly wounded tissue is 

inoculated. The spores are dispersed after fruiting bodies have absorbed sufficient moisture. Asexual spores 

(conidia) ooze out of saturated fruiting bodies and are 

dispersed by rainsplash and blown by wind. Occasionally 

insects or birds transport conidia. Sexual spores are 

forcibly discharged from saturated fruiting bodies into air currents, but occasionally they 

will ooze out as intact asci and then are forcibly discharged. Cytospora species infect 

senescing fruit spurs, shaded twigs, and senescing buds and bud scars. The fungus becomes an established and symptomless endophytic latent pathogen in scattered locations 

in the living wood, often residing in branch crotches. Trees with the latent pathogen may 

never exhibit disease or produce fruiting bodies on senescing tissues. The absence of the 

pathogen fruiting bodies in senescing tissues is due in part to rapid drying of the cambium 

tissues in climates like that of the Great Plains. Fruiting bodies are seldom evident in shed 

or broken branches when branch death or fall occurs during dry winter weather. Actively 

growing nonstressed trees limit expansion of Cytospora canker by their internal defense 

processes. Therefore, annual cankers occur on some trees, but perennial cankers occur on 

trees that do not have the genetic capability to limit canker expansion. Commercial peach 

cultivars have lost the capability of limiting canker expansion and have highly destructive 

perennial cankers, whereas other Prunus species generally retain tolerance and have annual cankers. 

Figure 25-4—Orange spore 

masses and tendrils above 

erumpent Cytospora pycnidia 

(William Jacobi, Colorado State 

University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 93

Fruiting bodies form in infected bark to complete the life cycle. The fungus overwinters as fruiting bodies and mycelium in bark. Several studies have concluded that despite 

the production of immense numbers of conidia on cankers, rarely produced ascospores 

are responsible for most infections. This disease cycle has been suggested for peach 

perennial canker indirectly through the use of population genetic markers, and was 

operating in orchards in which the sexual fruiting bodies were not found despite careful 

searching.

Damage

Cytospora canker can cause dieback or death of planted or native trees. It is a limiting 

factor in the establishment and growth of some trees. In nurseries, it can attack cuttings 

used for propagation of almost any hardwood, including poplars and willows. A disease 

called blackstem of cottonwood 

is caused primarily by C. chrysosperma (sexual stage Valsa 

sordida) and other canker fungi. 

It can develop during storage or 

after outplanting (fig. 25-5) and 

can result in severely reduced 

nursery production.

Management

Healthy trees are less susceptible to infection and damage. 

Drought stress is one of the most 

common predisposing factors 

with this disease. If possible, water and fertilize trees as needed to 

maintain optimum vigor. Reduce 

chances of infection by preventing wounds. Do not bring infected plant materials into 

the area. Plant resistant varieties if they are known or alternative species if they are acceptable. Severely cankered branches or trees should be removed and destroyed. Yearly, 

prune cankered branches back to a live branch beyond the canker. Small cankers on stems 

can be removed by excising all affected bark. Shape wounds into an ellipse to promote 

rapid healing. Prune during dry weather when rain or irrigation will not occur for two 

weeks (for example, in winter). Disinfect pruning tools with alcohol after each cut.

Fungicides are not normally effective in preventing infection or canker enlargement 

after infection.

To avoid cankers on nursery cuttings, maintain healthy stock blocks (mother blocks) 

for selecting your propagation wood. Collect shoots from stock blocks in the fall before 

very cold weather causes bark moisture to fall. Process and store cuttings quickly: dip 

cuttings in a fungicide solution before winter storage, store at a constant temperature 

below 35 °F, and plant cuttings in beds after soil warms to allow rapid plant growth.

Selected References

Kepley, J.B.; Jacobi, W.R. 2000. Pathogenicity of Cytospora fungi on six hardwood 

species. Journal of Arboriculture. 26: 326–333.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Walla, J.A.; Stack, R.W. 1980. Dip treatment for control of blackstem on Populus

cuttings. Plant Disease. 64: 1092–1095.

Figure 25-5—Development of blackstem symptoms following outplanting of poplar cuttings in a nursery setting 

(James A. Walla, Northern Tree Specialties, used with 

pemission).
94 USDA Forest Service RMRS-GTR-335. 2016.

26. Fire Blight of Apple, Pear, and Other Ornamental 

Rosaceous Shrubs and Trees

Megan M. Kennelly and Mark L. Gleason

Fire blight is caused by the bacterium Erwinia amylovora. The pathogen infects flowers, fruits, shoots, branches, and roots, leading to a wide array of symptoms. Flowers, 

fruits, shoots, branches, and even entire trees can be killed. Fire blight damages susceptible ornamental trees and shrubs and is a major threat to apple and pear production.

Hosts and Distribution

Fire blight occurs on many ornamental trees and shrubs in the family Rosaceae. This 

disease was first reported in New York in the 1700s and is now found throughout the 

United States. As of 2012 it had been reported in 47 countries in North America, Europe, 

the Mediterranean region, and the Pacific Rim. The fire blight pathogen is a quarantine 

organism in several important apple- and pear-producing countries. In the Great Plains, 

fire blight is common on apples and crabapples (Malus spp.) as well as fruiting and 

ornamental pears (Pyrus spp). Fire blight also occurs on cotoneaster (Cotoneaster spp.), 

hawthorn (Crataegus spp.), serviceberry (Amelanchier spp.), mountain-ash (Sorbus spp.), 

firethorn (Pyracantha spp.), and quince (Cydonia oblonga).

Symptoms and Signs

Infected flowers exhibit a water-soaked appearance, droop, then turn black or dark 

brown (fig. 26-1). Infections commonly begin with the flowers, and the pathogen then 

spreads into the shoot and leaves. Infected leaves and shoots turn dark brown or black, 

and shoot tips curl downward into a characteristic “shepherd’s crook” shape (fig. 26-2). 

Shorter side branches (spur branches) can also become infected (fig. 26-3). The blackened dead leaves and flowers remain 

attached, giving a scorched appearance to the tree. Gummy 

bacterial ooze may be present on shoots or fruit. Cankers can 

develop on branches of any size as well as the trunk (figs. 

26-4, 26-5, and 26-6). The bark on cankered areas is dark and 

may be sunken or cracked. Canker margins may be rough or 

smooth. When disease is active, cankers may have a watersoaked appearance with bacterial ooze present (figs. 26-4 and 

26-5). Tissue under the bark exhibits reddish-brown streaking 

(fig. 26-6). Branches with cankers are girdled and often die.

Figure 26-1—Flowers infected with fire blight. 

They droop and turn black or dark brown (Penn 

State Plant Pathology Collection, used with 

permission).

Figure 26-2—Fire blight-infected shoot with 

characteristic “shepherd’s crook” symptom 

(Megan M. Kennelly, Kansas State University, 

used with permission).

Figure 26-3—Fire blightinfected spur branch with 

blackened, “scorched” 

leaves (Megan M. Kennelly, 

Kansas State University, 

used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 95

Disease Cycle

Fire blight is most severe during 

moderately warm temperatures 

with rainfall. The optimum temperature range for infection is 70 

to 81 °F, with a minimum of 65 

°F and maximum of 90 to 95 °F. 

E. amylovora overwinters at the 

margins of cankers. During warm, 

wet weather in spring, the bacteria 

multiply, often dripping in a sticky 

ooze at canker margins. Bacteria are 

spread by rain, wind, and insects 

(primarily bees and flies) to cause 

primary infection on open flowers. 

The pathogen multiplies rapidly on 

the stigmas, and the bacterial cells 

can spread down into the nectaries. 

Once initial flowers are colonized, 

pollinating insects transfer the 

pathogen to other blossoms, and the 

bacteria can spread from flowers 

to supporting branches. Bacteria 

also infect the plant through natural 

openings and wounds in leaves and 

succulent shoots. These secondary 

cycles can occur throughout the 

growing season as long as conducive weather occurs and susceptible 

tissue is available. Wounding from 

wind and hail increases the risk of 

infection. Once inside the plant, 

bacteria can spread into larger 

branches, trunks, and roots. Root infection generally causes tree decline 

and death. 

Damage

Fire blight is a serious disease 

of commercial pome fruit in the 

United States and many other countries worldwide, reducing yields 

as well as directly killing entire 

branches and trees. Fire blight 

damage in ornamental trees and 

shrubs reduces ornamental value 

and functionality, with potential 

management and replacement costs. 

Disease severity varies from year to 

year based on weather conditions.

Figure 26-6—Small branch canker from fire blight with 

reddish-brown streaking under the bark (Penn State 

Plant Pathology Collection, used with permission).

Figure 26-4—Canker from fire blight exhibiting discoloration, cracking, and gummy bacterial ooze (George 

Sundin, Michigan State University, used with permission).

Figure 26-5—Canker from fire blight exhibiting bacterial 

ooze (Patricia McManus, University of Wisconsin – 

Madison, used with permission).
96 USDA Forest Service RMRS-GTR-335. 2016.

Management

There are several practices that should be used in combination, if possible, to prevent 

and manage fire blight. The following practices are focused on ornamental plantings. 

Detailed information on fire blight prevention in intensively managed commercial 

orchards is available from other sources and can be obtained from local Cooperative 

Extension personnel.Resistant varieties are available for some fruiting and flowering species, with varying levels of resistance. Consider using resistant varieties in new plantings. 

Cultural practices are also important to reduce the risk of fire blight. Succulent growth 

is more susceptible to fire blight. Therefore, maintain plants with appropriate irrigation, 

fertilization, and pruning to avoid overstimulation of growth. Several references listed 

below provide useful guidelines on cultivar selection and maintenance practices.

 If fire blight occurs, sanitation is critical to reduce further spread. The optimal time 

for pruning is during the dormant period, from mid- to late winter, and pruning cuts 

should be made at least 12 inches below any visible symptoms. If pruning in summer, 

however, conduct pruning during dry weather and make cuts at least 12 to 18 inches 

below any visible symptoms. In addition, wait to prune until late summer when new 

growth has ceased, as new succulent tissue is particularly susceptible. Disinfect pruning 

tools with a 1:10 bleach dilution between cuts. Pruned-out branches should be burned or 

discarded, not left at the site. Remove heavily infected trees.

Chemicals (primarily antibiotics and copper compounds), growth regulators, and 

biological controls are available to prevent fire blight development. If a tree has a history of fire blight, copper products can be applied before budbreak to reduce overwintering 

inoculum. Avoid applying copper once new growth begins, as phytotoxic reactions 

can occur, and follow all label instructions. Certain antibiotics (such as streptomycin 

products) are labeled for application during bloom to prevent blossom blight. Follow all 

label instructions to reduce the risk of antibiotic resistance, such as limits on number of 

applications per year and avoiding applications after symptoms have developed. Strains 

of E. amylovora with resistance to antibiotics have been reported in pome fruit orchards 

in several states. Exact details on the use of chemicals and biological controls are beyond 

the scope of this publication. Be sure to read, understand, and follow all label instructions 

for any pesticide including biological controls or growth regulators. Further information 

on fire blight management for commercial and home production of fruiting apple and 

pear is available in other references (see list below) and resources. Disease forecasting 

models to optimize the timing of applications are available for commercial apple and pear 

producers.

Selected References

Bell, A.C.; Ranney, T.G.; Eaker, T.A.; Sutton, T.B. Evaluating fire blight resistance 

among flowering crabapples (Malus spp.) and pears (Pyrus spp.). http://www.ces.ncsu.

edu/fletcher/programs/nursery/metria/metria11/bell/. (August 2014).

Gao, G.; Becker R.; Brown, M.; Ellis, M.; Prochaska, S. 2009. Midwest home fruit 

production guide: cultural practices and pest management. Bulletin 940. Columbus, 

OH: Ohio State University. 148 p.

Gilman, E.F. 2011. An illustrated guide to pruning. Cengage learning. 496 p.

Griffin, J. 2005. Flowering crabapples. Publ. MF-875. Manhattan, KS: Kansas State 

University Agricultural Experiment Station and Cooperative Extension Service. 8 p.

Hartman, J.R.; Pirone, T.P.; Sall, M.A. 2000. Pirone’s tree maintenance. 7th ed. New 

York: Oxford University Press. 560 p.

Johnson, D.T., et al. 2015. 2-15 Midwest Tree Fruit Spray Guide. Bulletin S-146, Kansas 

State University Agricultural Experiment Station and Cooperative Extension Service. 

68 p.

van der Zwet, T.; Orolaza-Halbrendt, N.; Zeller W. 2012. Fire blight: history, biology, and 

management. St. Paul, MN: APS Press. 460 p.

Vanneste, J.L. 2000. Fire blight: the disease and its causative agent, Erwinia amylovora. 

Wallingford, UK: CABI Publishing. 388 p.
USDA Forest Service RMRS-GTR-335. 2016. 97

27. Hypoxylon Canker of Aspen

Michael E. Ostry and Jared M. LeBoldus; revised from 

Thomas E. Hinds and Mark O. Harrell (Riffle and Peterson 1986)

Quaking aspen (Populus tremuloides) is the most widely distributed forest tree in 

North America, abundant throughout the northern boreal forest of Canada and extending as far south as Mexico. The tree is found in the northern states from Maine to 

Washington, but its occurrence in the Great Plains is sporadic, with small stands in 

Nebraska, North Dakota, and South Dakota. Fast-growing poplar hybrids are also commonly planted in the Great Plains. These natural and planted stands of Populus spp. are 

susceptible to infection by Hypoxylon canker.

Hosts and Distribution

Hypoxylon canker, caused by the fungus Entoleuca mammata (syn. Hypoxylon mammatum), is one of the most damaging diseases of quaking aspen in many areas of North 

America. Bigtooth aspen (P. grandidentata) is occasionally infected, as are hybrids of 

various Populus species. However, the disease is not nearly as damaging to these species 

as it is to quaking aspen, its most susceptible host.

Hypoxylon canker of aspen is abundant east of the Rocky Mountains and is a major 

cause of mortality of quaking aspen in some Great Lakes States (Minnesota, Wisconsin, 

and Michigan) and in the Northeast. Hypoxylon canker is of far less importance on aspen 

in its western range and has not been found in Alaska. In Canada the disease is widely 

distributed and in Europe Hypoxylon canker affects P. tremula. Although the disease is 

present in aspen stands on the northern Great Plains, its ecological importance in these 

scattered stands is unknown.

Symptoms and Signs

Cankers can develop anywhere on branches and stems of aspen of all ages, and 

symptoms vary depending on the stage of disease development. Dead leaves (“flags”) 

remaining on branches and twigs killed by the disease in the upper crowns are especially 

visible during the dormant season and often a symptom of the disease. Young cankers 

are slightly sunken, yellowish-orange areas with irregular margins (fig. 27-1). Later, the 

bark becomes blistered and ruptures, exposing the powdery gray mat of fungal tissue 

with bristlelike conidial pillars (hyphal pegs) (fig. 27-2), which bear single-celled, clear 

(hyaline) spores measuring 1.5 to 4.0 μm × 5.5 to 8.0 μm. 

Cankers become rough and black as patches of bark 

flake off. In the oldest areas of the cankers, fruiting 

bodies (perithecia) develop, embedded in hard, cushionlike stromata. Stromata are white when young and 

turn gray to black as they age (fig. 27-3). Within the 

fruiting bodies dark brown single-celled sexual spores 

Figure 27-1—A young Hypoxylon 

canker with sunken, yellowishorange bark and irregular margins 

(Michael E. Ostry, U.S. Forest 

Service).

Figure 27-2—Hyphal pegs beneath blistered bark of a young 

Hypoxylon canker (Michael E. Ostry, U.S. Forest Service).
98 USDA Forest Service RMRS-GTR-335. 2016.

(ascospores) measuring 9.0 to 12.0 μm × 

20.0 to 33.0 μm develop (fig. 27-4). 

Advancing canker margins are yellowish orange and often slightly sunken. 

Trees only occasionally are able to develop 

callus and subsequent woundwood to limit 

growth of the fungus, and most infections 

result in the characteristic diffuse-type 

canker. Cutting into cankers reveals a 

mottled black and yellowish-cream color 

characteristic of diseased sapwood, often 

with white or gray mycelial fans of the 

fungus extending well beyond the visible 

canker margin (fig. 27-5). Holes and areas 

where the bark has been removed at the 

margins of cankers of all ages are common. 

This damage is the result of birds searching 

for and excavating insect larvae of woodboring insects frequently associated with 

the diseased wood. 

Disease Cycle

Aspen bark contains chemicals toxic to 

the fungus, so a wound through the bark 

and into the xylem is needed for infection. 

Oviposition wounds on branches made by 

several species of insects are commonly 

the sites of infection. Birds and insects 

could also inadvertently introduce bits 

of diseased tissues or mycelium of the 

fungus into wounds during their foraging 

activities. The fungus colonizes and obtains 

energy from the sapwood to advance under 

the bark beyond the wound and produces a 

toxin that inhibits wound callus formation. 

This toxin kills the bark, resulting in canker 

development. The fungus can have a latent 

period of two years or more before symptoms develop.

Approximately one year after initial 

symptoms appear, the hyphal pegs develop 

and rupture the bark, exposing hyaline spores that are not infective but rather function 

as gametes playing a role in the production of perithecia one to two years later. Singlecelled, dark brown ascospores are released from perithecia in the cushionlike stromata 

during wet weather and infect trees through various wounds, completing the life cycle of 

the fungus. 

Damage

Hypoxylon canker girdles and kills twigs, branches, and stems of trees. Disease 

incidence and impact are the greatest in the first 20 years of a developing aspen stand. 

Mortality of trees of this age is common because cankers are lower on the stem than on 

older trees. Older trees can recover from stem cankers higher in their crowns by developing new leaders. Wind breakage of weakened stems at cankers is common owing to the 

wood decay that develops (fig. 27-6). Aspen in plantations and ornamental landscapes 

are especially vulnerable to damage by Hypoxylon canker because infection of persistent 

lower branches on widely spaced trees often results in stem canker development.

Figure 27-3—A Hypoxylon canker with rough, 

blackened bark and gray perithecial stromata 

(Michael E. Ostry, U.S. Forest Service). 

Figure 27-4—Sexual spores (ascospores) 

of Entoleuca mammata. Scale bar is 10 μm 

(Michael E. Ostry, U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 99

Figure 27-6—Stem breakage at a Hypoxylon canker (U.S. Forest Service).

Management

There are no direct control measures 

proven to minimize the impact of 

Hypoxylon canker over a large area. 

Susceptibility to the disease varies by 

clone. Therefore, favoring any resistant 

clones identified in native stands or selecting and planting genetically resistant trees 

have been suggested as ways to reduce 

damage in natural stands and plantations.

Hypoxylon canker frequently infects 

trees through wounds on branches made by 

various insects that are common in aspen 

along stand edges, in understocked stands, 

and in plantations with widely spaced 

trees. As a result, maintaining fully stocked 

stands and avoiding openings are recommended control strategies. Self-pruning of 

lower branches that often are infected and 

result in lethal stem cankers is enhanced by 

shading in dense stands and has been found 

to be a genetic trait of canker-resistant 

aspen in test plantings. Individual trees 

may be saved by early pruning of diseased 

branches before the fungus enters the main 

stem.

Figure 27-5—Mottled black and yellowish-white 

sapwood and white mycelial fan beneath a 

Hypoxylon canker (gray perithecial stromata, 

U.S. Forest Service).

Selected References

Ostry, M.E.; Anderson, N.A. 1998. Interactions of insects, woodpeckers, and Hypoxylon 

canker on aspen. Res. Pap. NC-331. St. Paul, MN: U.S. Department of Agriculture, 

Forest Service, North Central Research Station. 15 p.

Ostry, M.E.; Anderson, N.A. 2009. Genetics and ecology of the Entoleuca mammataPopulus pathosystem: implications for aspen improvement and management. Forest 

Ecology and Management. 257: 390–400.
100 USDA Forest Service RMRS-GTR-335. 2016.

28. Phomopsis Canker of Poplar

Michael E. Ostry and Jared M. LeBoldus; revised from: Theodore H. 

Filer Jr. and Edward M. Sharon (Riffle and Peterson 1986)

Cottonwood (Populus spp.) occurs along streams and rivers throughout the Great 

Plains. In most of its range, cottonwood grows on moist, well-drained soils; but in the 

western semi-arid one-third of its range, it is found only along streams. Most species of 

Populus and their interspecific hybrids are susceptible to Phomopsis canker.

Hosts and Distribution

Phomopsis species cause dieback and cankers on many tree species in North America, 

as well as in Europe and Japan. Phomopsis macrospora commonly affects Populus 

species in nurseries and young plantations. The 

teleomorph of the fungus has not been reported in 

North America. Outbreaks of Phomopsis canker 

have been reported from Minnesota to Mississippi 

and in Japan. 

Symptoms and Signs

Cankers caused by Phomopsis species develop 

on cottonwood twigs, limbs, and boles; they are 

inconspicuous during early development (fig. 28-1). 

Small pustules (pycnidia) on cankers appear as small 

black dots protruding through bark epidermis (fig. 

28-2). Phomopsis fruits profusely on dead tissues 

(fig. 28-3). Spores are pushed out of the pore of 

the pustules under moist conditions and often form 

sticky white, orange, or reddish spore tendrils (fig. 

28-4). Two types of spores are produced: spindleshaped alpha spores that are 11.0 to 20.5 μm × 2.5 to 

5.0 μm and J-shaped beta spores 10 to 20 μm × 1.3 

to 2.5 μm (fig. 28-5).

As the canker develops and 

enlarges, the phloem tissue 

dies, forming a sunken area. 

When the stem is girdled, the 

distal portion dies; the resulting dead leaves cling to the 

branches for several weeks. 

Similar symptoms are caused 

by Fusarium, Cytospora, and 

other canker-causing fungi.

Disease Cycle

The asexual spores 

(conidia) are the primary 

inoculum. Conidia can be 

dispersed by wind and insects, 

and rainsplash often spreads 

the spores to wounds on other 

parts of the tree and to adjacent trees. The fungus is considered a wound parasite, and requires a wound or natural 

opening before it can penetrate and colonize tissues. The size of the canker that develops 

depends on tree vigor. On fast-growing cottonwoods, canker development is usually 

limited by woundwood development, which restricts fungal growth. On slow-growing 

Figure 28-1—Phomopsis stem 

canker on hybrid poplar (Michael E. 

Ostry, U.S. Forest Service).

Figure 28-2—Poplar cuttings affected by blackstem disease 

caused by Phomopsis macrospora during improper storage, 

with small black fruiting bodies (pycnidia) protruding through 

the bark (Michael E. Ostry, U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 101

Figure 28-3—Phomopsis canker 

on planted cutting (Michael E. 

Ostry, U.S. Forest Service).

Figure 28-4—Spore tendrils of Phomopsis macrospora on diseased cutting (Michael E. Ostry, U.S. Forest Service).

Figure 28-5—Spindle-shaped 

alpha spores and J-shaped beta 

spores of Phomopsis spp. Scale 

bar is 20 μm (Michael E. Ostry, 

U.S. Forest Service).

trees (trees growing under moisture stress or in poorly 

aerated soils), cankers can girdle the stem and cause 

tree mortality.

Damage

Phomopsis is one of the most important pathogens 

that cause mortality of water-stressed planted cuttings. 

In most cases the fungus is on or in the bark of cuttings 

when they are harvested from the nursery. If cuttings 

are under water stress the fungus can girdle the cutting 

within a few weeks of planting.

Management

Good cultural practices can reduce the likelihood 

of mortality of shade trees. Water at the rate of two 

inches per week during periods of moisture deficit, and 

control insect defoliators and leaf diseases to reduce 

tree stress and promote tree vigor. Fertilize prudently, 

applying needed nutrients only if a soil test shows a 

deficit.

To prevent losses when planting poplars, select 

healthy, canker-free cuttings or seedlings. Prevent 

cuttings from drying in storage; soak them for 48 hours in water to initiate root primordia 

before planting. Cultivation is needed during the first year to keep trees growing vigorously. Plant poplar clones that are resistant to other diseases whenever possible.

Selected References

Ostry, M.E.; McNabb, H.S., Jr. 1982. How to identify and prevent injury to poplars 

caused by Cytospora, Phomopsis, and Dothichiza. HT-57. St. Paul, MN: U.S. 

Department of Agriculture, Forest Service, North Central Forest Experiment Station. 

6 p.
102 USDA Forest Service RMRS-GTR-335. 2016.

29. Septoria Leaf Spot and Canker of Poplar

Jared M. LeBoldus, Michael E. Ostry, and James T. Blodgett; revised 

from Joseph M. Krupinsky and David W. Johnson; Jerry W. Riffle 

and David S. Wysong (Riffle and Peterson 1986)

Sphaerulina musiva (syn. Septoria musiva) causes a leaf spot disease and branch and 

stem cankers on native and hybrid poplars (Populus spp.). The sexual form of this fungus 

is Mycosphaerella populorum. There are two closely related species, S. populicola and 

M. populi, which also cause a leaf spot disease of poplar, as well as M. jaczewskii and 

S. aceris, which infect caragana (Caragana spp.) and maple (Acer spp.), respectively.

Hosts and Distribution

S. musiva is indigenous throughout much of the United States and Canada and occurs 

across the Great Plains. This fungus causes leaf spots on native cottonwood (P. deltoides) 

and hybrid poplars. It also causes a branch and stem canker disease of hybrid poplars 

and introduced Populus species. In the North Central region of the United States hybrids 

among cottonwood, balsam poplar (P. balsamifera), black poplar (P. nigra), Japanese 

poplar (P. maximowiczii), black cottonwood (P. trichocarpa), and laurel poplar (P. laurifolia) are common hosts.

Symptoms and Signs

Leaf spot symptoms vary according to time of infection, host species, and age of 

leaves. Four types of leaf spot symptoms have been described: (a) brown, mostly circular leaf spots (1 cm in diameter) that may have a brown or yellow margin, with black, 

pimplelike fruiting bodies (pycnidia) clustered within; (b) small flecks, commonly with 

very angular margins; (c) white or silvery spots, mostly 1 to 3 mm in diameter; and 

(d) irregularly shaped spots that are light tan in the center with brown margins (fig. 29-1). 

Numerous black pycnidia are commonly clustered in the center of these large spots. 

Immature pycnidia are light brown and turn black when mature (fig. 29-2).

Cankers are formed on the main stem and branches of the current season’s growth. 

Cankers are often flat-faced or have swollen marginal callus and early woundwood 

(fig. 29-3). The bark over young cankers is dark brown or black (fig. 29-4) and depressed. 

Figure 29-1—Leaf spot symptoms on 

three balsam poplar leaves (Jared M. 

LeBoldus, Oregon State University, used 

with permission).

Figure 29-2—Immature asexual fruiting bodies (pycnidia) 

clustered in the center of a leaf spot (Jared M. LeBoldus, 

Oregon State University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 103

Infected cambium is killed and small 

black pycnidia may develop in the 

bark on the ashy-white central area 

of cankers. Continued development 

of cankers may result in girdling and 

death of affected branches and stems 

during late summer. Affected stems 

may be infected by other canker fungi, 

such as Cytospora chrysosperma, 

resulting in additional damage.

Two types of spores develop 

within fruiting bodies in infected 

host tissues. Asexual spores (conidia) 

develop in pycnidia on cankers and 

leaf spots throughout the growing 

season, and are exuded in pink or 

white tendrils during wet weather. 

Conidia are clear, cylindrical, straight 

or curved, and one- to four-septate, 

and measure 20 to 56 μm × 3 to 4 μm 

Figure 29-4—Canker (necrotic lesion) caused by Sphaerulina musiva on a young poplar shoot 

(Jared M. LeBoldus, Oregon State University, used with permission).

Figure 29-3—Multiple cankers caused by Sphaerulina 

musiva on a seven-year-old balsam poplar stem 

with a broken top (Jared M. LeBoldus, Oregon State 

University, used with permission).

(fig. 29-5). Sexual spores (ascospores) are produced in perithecia that develop on fallen, 

infected leaves. The ascospores are clear, one-septate, and 16 to 28 μm × 4.5 to 6.0 μm. 

Fungi other than M. populorum can cause similar leaf spots (listed above). Microscopic 

examination of spores or a molecular technique (i.e, internal transcribed spacer [ITS] 

sequences) is necessary to identify these pathogens.

Disease Cycle

The pathogen overwinters on fallen infected leaves and in branch cankers. In spring, 

ascospores from fallen leaves and conidia from cankers are discharged during wet 

weather. These spores are dispersed by wind and rainsplash to infect newly emerging leaf 

and stem tissue. Infections may occur through stipules, petioles, buds, lenticels, or bark 
104 USDA Forest Service RMRS-GTR-335. 2016.

wounds. Leaf infection usually precedes stem infection. Leaf spots develop soon after 

leaf emergence and the fungus spreads to stems and branches, causing cankers. Cankers 

begin as necrotic lesions on the current season’s growth. Pycnidia usually develop shortly 

after and produce conidia that are dispersed and cause secondary infections. Disease 

development is enhanced by warm temperatures and long periods of high humidity.

Damage

S. musiva damages poplars of all ages, but damage is most severe in nursery environments, young plantations, and windbreaks. Numerous leaf spot infections can result 

in premature defoliation of susceptible clones. Multiple cankers can girdle stems, and 

affected trees are susceptible to wind breakage. Cankers are infection courts for other 

pathogens such as Cytospora, Phomopsis, and Fusarium and decay fungi, all of which 

can contribute to the damage. 

Management

Damage caused by S. musiva is reduced primarily by the use of resistant cultivars. 

Clones resistant or moderately resistant to leaf spot may also be more resistant to stem 

canker and should be selected for planting. Use vigorous, disease-free planting stock for 

maximum early growth and to avoid introducing pathogens into new plantings.

Cultural treatments, such as cultivation or raking in the fall to remove leaf litter 

harboring fungal inoculum, will minimize primary infection in spring if inoculum from 

adjacent trees is not a factor. Planting moderately susceptible trees at a wide spacing to 

provide good air circulation within the canopy will reduce the duration of free moisture 

on leaves and minimize infection by S. musiva. 

Selected References

LeBoldus, J.M.; Blenis, P.V.; Thomas, B.R. 2008. Clone by isolate interaction in the 

hybrid poplar-Septoria musiva pathosystem. Canadian Journal of Forest Research. 38: 

1888–1896.

LeBoldus, J.M.; Blenis, P.V.; Thomas, B.R.; Feau, N.; Bernier, L. 2009. Susceptibility 

of Populus balsamifera to Septoria musiva: a field study and greenhouse experiment. 

Plant Disease. 93: 1146–1150.

Newcombe, G.; Ostry, M. 2001. Recessive resistance to Septoria stem canker of hybrid 

poplar. Phytopathology. 91: 1081–1084.

Ostry, M.E.; McNabb, H.S., Jr. 1986. Populus species and clones resistant to 

Melampsora, Marssonina, and Septoria. Res. Pap. NC-272, U.S. Department of 

Agriculture, Forest Service, North Central Forest Experiment Station. 7 p.

Figure 29-5—Asexual spores (conidia) of Sphaerulina musiva. Scale bar 

measures 20 μm (Jared M. LeBoldus, Oregon State University, used with 

permission).
USDA Forest Service RMRS-GTR-335. 2016. 105

30. Stem Decays

James J. Worrall

Stem decay is the decay of wood in aboveground stems of live trees. Fungi that cause 

stem decays in the Great Plains occur in several orders of phylum Basidiomycota, class 

Agaricomycetes. Almost all are polypores, with fruiting bodies that are often called 

conks. 

Hosts and Distribution

Stem decays occur in all tree species throughout the Great Plains, but tree species 

differ greatly in the frequency and severity of decays. Decays also may vary in frequency 

over the distribution of the host. 

Symptoms and Signs

Conks are often the first outward evidence of infection (fig. 30-1), but they only appear years after decay begins and may never appear, even when the tree is extensively 

decayed. In advanced cases, decay may be detected by sounding: hitting the bark with 

a mallet or back of a hatchet and listening for a somewhat hollow sound. Often, decay 

is exposed in broken stems, branch stubs, split forks, or other openings. Otherwise, the 

only recourse for diagnosis is cutting the 

tree open or using an increment borer to 

sample the wood.

Crown symptoms are not common with 

stem decays, but can occur. Some stem 

decay fungi attack the sapwood and can 

even cause cankers. Crown thinning and 

dieback of branches often result. 

Fungi that attack live trees may continue to decay in and fruit on the tree after 

it dies. Other fungi grow mostly on dead 

trees; some common representatives are 

included here. It is important to recognize 

that fungi fruiting on dead trees or dead 

portions of trees may not have decayed 

wood or killed tissues in the live tree.

Two major types of wood decay are 

caused by the fungi considered here. In uniform white rot, all structural components 

(cellulose, hemicelluloses, lignin) are decomposed, and partly decayed wood has about 

the same proportion of components as undecayed wood. Wood loses strength and becomes soft, but retains its fibrous nature, often becoming stringy. Decayed wood leaves 

a fibrous, splintery fracture surface when it breaks. Usually white-rotted wood does not 

darken appreciably and may become bleached white.

In brown rot, cellulose and hemicelluloses are consumed, but lignin remains more 

or less intact. Cellulose is depolymerized early in the decay process, so that the fibrous 

nature of the wood is lost and wood breaks readily across the grain as well as with it. In 

advanced decay, the residual wood shrinks, giving a cubical appearance. Decayed wood 

leaves relatively nonsplintery, cross-grain, flat surfaces when it breaks (often called a 

brash failure). Wood with advanced brown rot crumbles readily to a powder when dry. 

The lignin becomes somewhat darker during the decay process, turning the residual wood 

brown.

Disease Cycle

Polypores are so named because the conk has on its lower surface thousands of small, 

vertical tubes, each of which ends at the bottom with a pore (fig. 30-2d, e). Spores 

(microscopic propagules) are produced in the tubes, fall through the pores, and are then 

wind dispersed. If a spore chances to land on a suitable point of infection under moist 

Figure 30-1—Conk of Fomes fomentarius on common horsechestnut (Aesculus 

hippocastanum) (Andrej Kunca, National Forest 

Centre—Slovakia, Bugwood.org).
106 USDA Forest Service RMRS-GTR-335. 2016.

conditions, it will germinate and the fungus will grow into the host. The infection site 

differs among fungi and is not well known in many cases. For many fungi, especially saprobic species, the infection sites generally are substantial wounds, such as large branch 

stubs or mechanical scars. More highly adapted parasites may infect smaller sites such as 

dead twigs or twig scars. In either case, microscopic, branching filaments (hyphae) grow 

into the wood and eventually cause decay. Over many years, the wood provides sufficient 

resources for fruiting. For most fungi, fruiting also requires that two individuals of the 

same species and compatible mating type encounter each other in the wood and fuse 

together. Fruiting completes the cycle. 

Figure 30-2—Fomitiporia texana on oneseed juniper (Juniperus monosperma) in Whitby Canyon, 

Comanche National Grassland, Colorado. All photos are from the same tree, which had eight conks. 

(a) Crown with estimated 90 percent dieback. (b) Four of the conks on the stem. (c) Closeup of the 

lowermost conk in B). (d) Pore surface. (e) Vertical section through a conk showing several layers 

of tubes and context at top left. (f) Vertical section of the stem at 3 to 6 ft above ground. The sapwood is extensively decayed on both sides, and an inner column of decay is connected to a branch. 

(g) Cross-section with conk about 6 ft above ground. Decay is much more extensive in the sapwood 

than in the heartwood, and only a slight amount of sapwood is left alive. (h) Uniform white rot with 

zone lines. (i) White stringy rot where fine whisps of brown mycelium may be observable (James J. 

Worrall, U.S. Forest Service. Location and dissection provided by Jeremy Schoonover, U.S. Forest 

Service).
USDA Forest Service RMRS-GTR-335. 2016. 107

Damage

Decay in stems and branches increases the likelihood of mechanical failure, resulting 

in loss of portions of the crown or sometimes the entire live crown. Assuming a concentric decay column, likelihood of failure increases substantially when less than one-third 

of the stem radius remains sound.

Such mechanical failure can be hazardous if people or their property are nearby. In 

developed sites (around buildings, roads, points of interest, and recreation areas, where 

people are concentrated), trees should be managed to assess such hazard and mitigate it as 

needed. Recognizing indicators of decay, understanding how decay develops, and identifying fungi as parasitic versus saprobic can all be important in assessing trees for hazard.

Incidence of decay is generally correlated with tree age. Young trees usually are 

free of decay. As trees age, branch stubs, wounds, and time increase the opportunity for 

infection. Once decay is established and begins expanding, time favors the fungus. Older 

trees are less vigorous, have a greater proportion of nonliving heartwood, and grow more 

slowly. They are thus less resistant to decay pathogens, have more resources available for 

fungi that become established in the heartwood, and are less likely to keep up with decay 

by adding wood faster than it is lost.

Many windbreaks in the Great Plains were established during the Prairie States 

Forestry Project, 1935-1942. For most species, these have reached an age of declining 

vigor and increased damage from stem decays.

Buffaloberry (Shepherdia spp.) can be severely damaged by Perenniporia ellisiana in 

the northern Great Plains. This fungus may destroy windbreak plantings that are only 20 

to 25 years old. Buffaloberry is so susceptible that its extensive use in the northern Great 

Plains is discouraged. 

The most common stem decay of black locust (Robinia pseudoacacia) is caused by 

Fulvifomes robiniae, which is host-specific. The disease is common in the central and 

southern Great Plains, particularly in Oklahoma, where about 25 percent of surveyed 

black locusts in 40-year-old windbreaks were infected. As with stem decays in general, 

incidence increases with tree age. 

Fomitiporia punctata attacks many hardwoods and is common in the northern Great 

Plains, particularly in North Dakota. There, the disease occurred in about one-third of 

sites with trees more than 20 years old. Based on incidence of conks, F. punctata was 

the most important stem decayer of green ash (Fraxinus pennsylvanica) in Prairie States 

Forestry Project windbreaks in North Dakota. It tends to cause a canker-rot, both decaying the inner wood and growing to the surface and killing patches of sapwood and bark. 

The most common stem decay of ashes in the Great Plains is caused by Perenniporia 

fraxinophila (fig. 30-3). This fungus occurs on other species as well, including bur oak 

(Quercus macrocarpa) and boxelder (Acer negundo). 

Green ash in woodlands and windbreaks of the central 

and northern Great Plains is often heavily infected. In 

native woodlands of Nebraska, more than 10 percent 

of trees were estimated to be infected. A consistent 

straight-line relationship was found between diameter 

and percent infection (fig. 30-4). Conks were less frequent in a large sample of windbreaks 35 to 42 years 

old in Nebraska, occurring in 90 percent of windbreaks and 5.5 percent of trees. A contemporary study 

in North Dakota found only 47 percent of windbreaks 

and 0.3 percent of trees with conks. Of course, these 

estimates are conservative because additional trees 

were infected but without conks. 

More recently, in woodlands of east-central 

Montana, P. fraxinophila was associated with widespread canopy dieback of green ash. Stands had on average 38 percent incidence of 

conks; in some stands as many as 67 percent of trees had conks. Trees with conks had 

greater crown dieback than trees without conks. Evidence suggested that P. fraxinophila

is a cause of the widespread crown dieback, but is unlikely to be the sole cause.

Figure 30-3—Conks of 

Perenniporia fraxinophila (James 

J. Worrall, U.S. Forest Service).
108 USDA Forest Service RMRS-GTR-335. 2016.

Both F. punctata in North 

Dakota and P. fraxinophila in 

Nebraska windbreaks were found 

fruiting more commonly in the 

east than in the west. In Nebraska 

windbreaks, the difference was suggested to have been in part because 

the higher precipitation in the east 

promoted more fruiting and infection. In Nebraska woodlands, Riffle 

and others (1984) did not suggest 

any pattern in P. fraxinophila incidence across the state. However, 

Lesica and others (2003), reanalyzing Riffle and others’ (1984) 

data with their single datum from 

eastern Montana, suggested that the 

pattern was the opposite, and that 

drier areas had more fruiting.

Eastern redcedar (Juniperus 

virginiana), a major component of 

field windbreaks in the Great Plains, is frequently infected by Antrodia juniperina in the 

southern and central Great Plains. This pathogen may also attack other juniper species. 

The disease does not occur everywhere the host does, and its distribution is erratic. The 

fungus apparently enters through dead branches and wounds, then causes extensive decay 

in the stem, killing sapwood in the process. Stems become hollow and often break during 

storms. A windbreak may deteriorate if infected junipers are a major component.

As noted above, decay fungi can sometimes contribute to low vigor, thinning crowns, 

and branch dieback, to the extent that they kill sapwood from the inside. If trees are not 

able to produce new sapwood as fast as it is lost, they will suffer from the loss of water 

conduction and food storage. An example is decay of junipers by Fomitiporia texana (fig. 

30-2). This aggressive pathogen attacks the sapwood at an early stage, leading to crown 

thinning and dieback of branches. It eventually kills cambium and can kill the tree. Trees 

with a single conk may already have thin crowns, several conks are usually associated 

with dieback, and trees are often near death when the number of conks approaches 10. 

Taxonomy and Identification

Some basic terminology is helpful in identifying conks:

Context—the internal tissue of the conk above the tube layers (or for resupinate 

fungi, between the tube layers and substrate).

Effused—spread out over the substrate indeterminately.

Pileate—having at least a portion of the conk with a sterile upper surface over a fertile lower surface. This portion is called a pileus, although the term may be used 

more specifically in describing the upper surface.

Reflexed—mostly resupinate, but with a small portion at the upper end that is 

curved out, creating a small pileus.

Resupinate—lying back on the wood or bark, thus not having a pileus. Resupinate 

polypores generally show only a pore surface surrounded by a narrow margin.

Sessile—Pileate, without a stem, and not effused; typical conk form

Trama—the tissue between the tubes.

Figure 30-4—Percentage of green ash trees with conks 

of Perenniporia fraxinophila in relation to diameter at 

breast height (based on data from Riffle and others. 

[1984]).
USDA Forest Service RMRS-GTR-335. 2016. 109

Key to Major Wood Decay Fungi of the Great Plains

The key primarily uses field characters. However, a microscope and knowledge of 

polypore structure and anatomy will be needed in some groups and are helpful for confirmation of all species. Species characters are based primarily on Gilbertson and Ryvarden 

(1986), Lowe (1942, 1957), and Overholts (1953). Most species in the key are those 

included in Riffle and Peterson (1986); table 30-1 cross-references modern and 1986 

names.

1. Causing a brown rot; pileus (at least when young), pore surface and context white, 

cream, yellow, orange, pale brown or pale pink to rosy brown when fresh; hyphal 

system dimitic (if trimitic, then also with fusiform cystidioles); generative hyphae with 

clamp connections (if simple-septate, then accompanied by binding hyphae rather than 

skeletal hyphae); spores cylindrical or allantoid (if ovoid-ellipsoid, then generative 

hyphae simple-septate), hyaline, thin-walled . . . . . . . . . . . . . . . . . . Fomitopsidaceae

1. Causing a white rot; not with the above combination of characters . . . . . . . . . . . . . . 2

2. Pore surface when fresh white, cream, or yellowish buff, bruising to dark brown or 

purplish brown; conk perennial; context and tubes dark brown to purplish brown; 

spores with a truncate apex, ovoid, some broadly ellipsoid, brown, with a two-layered 

wall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ganodermataceae

2. Fresh pore surface not substantially changing color when bruised or bruising dull red 

and spores not as above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

3. Conk perennial, usually hard and woody; pileus dark yellowish to reddish brown or 

black; context dark yellowish or reddish brown, turning permanently black in potassium hydroxide (KOH); pore surface brown; pores circular and usually quite small 

with thick dissepiments; generative hyphae simple-septate; setae often present in the 

hymenium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hymenochaetaceae

3. Conk annual or perennial, tough and leathery to corky rather than woody; pileus, context, and pore surface usually white to pale brown; setae absent. If colors are similar to 

Hymenochaetaceae (such as in Fomes fomentarius), then with a distinct granular core 

in the context at the point of attachment .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

4. Conk annual (if perennial then tube layers not separated by layer of context); 

hyphal system dimitic to trimitic; hyphae with clamp connections; cystidia 

absent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Polyporaceae

4. Conk perennial, with distinctly stratified tube layers separated by layer of context; pileus (at least when young), pore surface, and context cream to buff; pores circular and 

small, 5–7 per mm; hyphal system monomitic; hyphae simple-septate; hymenium with 

encrusted cystidia. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Oxyporus populinus (fig. 30-7)

Fomitopsidaceae

1. When fresh, pileus yellow to orange; pore surface yellow to white; context white; 

generative hyphae simple-septate and accompanied by binding hyphae; spores ovoid to 

ellipsoid. . . . . . . . . . . . . . . . . . . . . . . . . . Laetiporus sulphureus sensu lato (fig. 30-8)

1. Colors different from above; generative hyphae with clamp connections and accompanied by skeletal hyphae; spores cylindrical or allantoid. . . . . . . . . . . . . . . . . . . . . . . . . 2

2. Conk resupinate to effused-reflexed (rarely sessile); white, cream or pale woody 

brown throughout (pileus may blacken with age); pores large (≤ 4 per mm), round and

regular but elongated and sinuous on sloping surfaces . . . . . . . . . . . . . . . . . . Antrodia

2. Conk sessile to effused-reflexed (rarely resupinate); colors pinkish to rosy brown or 

ivory/tan; pores smaller (≥ 4 per mm), round to angular . . . . . . . . . . . . . . . Fomitopsis

Antrodia

a. Typically on live junipers, especially eastern redcedar; conk effused-reflexed to 

nodulose; pileus pale buff to cork-colored, weathering gray to nearly black; context 

cream-colored; pore surface white; pores up to 1–3 mm wide and angular to daedaleoid; spores 6.5–9.0 μm x 2.5–3.5 μm . . . . . . . . . Antrodia juniperina (fig. 30-9)
110 USDA Forest Service RMRS-GTR-335. 2016.

a. Typically on dead hardwoods, especially willows; not with the above combination 

of characters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . b

b. Pileus and pore surface white to cream; pores 2–3 per mm on horizontal 

surfaces, but tending to become lamellate; spores 10–14 μm x 3.5–5.0 μm 

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antrodia albida (fig. 30-10)

b. Pileus and pore surface pale woody brown; pores 3–4 per mm on horizontal surfaces; spores 7–10 μm x 2.5–4.0 μm. . . . . . . . . . . . . . . . . . . . . Antrodia malicola

Fomitopsis

a. Conk perennial; pink to rosy brown throughout; pores 4–5 per mm; hyphal system 

dimitic; spores allantoid; uncommon . . . . . . . . . Fomitopsis cajanderi (fig. 30-11)

a. Conk annual or biennial; ivory to tan throughout; pores 5–7 per mm; hyphal system 

trimitic; spores cylindrical and slightly fusiform . . . . . . . . . . . . Fomitopsis meliae

Ganodermataceae

1. On various parts of live and dead hardwoods; conk perennial; crust on pileus usually 

thick and hard; pore surface white when fresh, quickly bruising dark brown, becoming 

dull buff when dry . . . . . . . . . . . . . . . . . . . . . . . Ganoderma applanatum (fig. 30-12)

1. On dead hardwoods; conk usually biennial; crust on pileus thin and easily indented; 

pore surface cream to yellowish when fresh, bruising or drying dull purplish brown 

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ganoderma lobatum

Hymenochaetaceae

1. On live shrubs, mostly currant (Ribes spp.); conk soft when fresh but hard when 

dry; pores almost invisible to naked eye, 6–7 per mm; hyphal system monomitic; 

spores pale yellow, subcylindrical; setae absent; widespread but uncommon in the 

Great Plains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phylloporia ribis

1. On live or dead trees; conk hard and woody; pores mostly larger than above . . . . . . 2

2. Typically on dead trees or dead portions of live trees; pileus not becoming rimose 

(deeply cracked, figs. 30-2c and 30-5); setae abundant; spores hyaline, ellipsoid to 

subglobose, nondextrinoid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2. On live trees; pileus becoming rimose; microscopic features variable. . . . . . . . . . . . . 4

3. Conk annual or short-lived, sessile or occasionally effused-reflexed; causing decay of 

dead or live trees, especially green ash, black locust, maple, and willow; setae arising 

from subhymenium; encrusted hyphae in hymenium or dissepiments; spores thinwalled, ellipsoid to ovoid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fuscoporia gilva

3. Conk perennial; sessile to resupinate; causing decay of primarily dead hardwoods; 

hyphae not encrusted; setae arising from tramal hyphae; spores somewhat thick-walled 

and ovoid to subglobose . . . . . . . . . . . . . . . . . . . . Phellinopsis conchata (fig. 30-13)

4. Spores hyaline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

4. Spores yellowish to reddish brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

5. Context usually dark reddish brown; hyphal system distinctly dimitic; spores not dextrinoid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phellinus

5. Context brass-colored (yellowish brown); hyphal system weakly dimitic; spores 

strongly dextrinoid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fomitiporia

6. Practically restricted to black locust; context light reddish brown; hyphal system 

dimitic; spores thin-walled, ellipsoid to ovoid, and flattened on one side 

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fulvifomes robiniae (fig. 30-14)

6. On oak, walnut, or occasionally other hosts; context dark reddish brown; spores thickwalled, ovoid to subglobose; hyphal system monomitic in context and dimitic in 

trama . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

7. On live walnut in the central and southern Great Plains; setae rare to frequent, 20–52 

μm x 6–14 μm; spores pale yellowish brown . . . . . . . Inonotus weirianus (fig. 30-6)

7. Typically on live oaks; setae frequent to abundant, 16–36 μm x 5–10 μm; spores distinctly reddish 

brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phellinus everhartii (fig. 30-5)
USDA Forest Service RMRS-GTR-335. 2016. 111

Fomitiporia

a. Conk sessile; on live junipers or hardwords; known only in the southern Great 

Plains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fomitiporia texana (fig. 30-2)

a. Conk resupinate; on live or dead hardwoods in most of the Great Plains 

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fomitiporia punctata (fig 30-15)

Phellinus

a. On live aspens; conk triangular in profile, upper and lower surfaces at approximately 45° from horizontal . . . . . . . . . . . . . . . . . . Phellinus tremulae (fig. 30-16)

a. On other hosts; pore surface orientation variable . . . . . . . . . . . . . . . . . . . . . . . . . . b

b. On live Prunus; pores 7–9 per mm . . . . . . . . . . . Phellinus pomaceus (fig 30-17)

b. On many live hardwoods; pores 5–6 per mm . . . . . Phellinus igniarius sensu lato 

(fig. 30-19)

Polyporaceae

1. Conk perennial; pores round; spores either large (12–18 μm long) or thick-walled, 

truncate, and dextrinoid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

1. Conk annual; pores round, angular, elongated, daedaleoid, or even lamellate; spores 

≤12 μm long (or if longer, pileus densely hirsute to hispid), thin-walled, neither truncate nor dextrinoid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2. Conk sessile, ungulate, woody; pileus gray to brown in sulcate zones, with a hard glabrous crust; pore surface concave, brown; context thin, dark yellowish brown, 

with a granular, mottled core at point of attachment; older tubes often stuffed with 

white mycelium; spores thin-walled, neither truncate nor dextrinoid, 12–18 μm x 

4–7 μm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fomes fomentarius (fig. 30-1)

2. Conk resupinate to sessile; pileus smooth, cream to black with age; pore surface flat 

or convex; pore surface and context white to light tan; spores thick-walled, truncate, 

dextrinoid. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Perenniporia

3. On dead hardwoods; conk resupinate to effused-reflexed; pileus dark brown to black; 

pore surface white to pale brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Datronia

3. On dead or live hardwoods; conk pileate and sessile; pileus and pore surface usually 

white to medium brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

4. Pileus finely tomentose to glabrous; pileus, context, and pore surface white to creamy 

buff when fresh . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Trametes

4. Pileus usually hirsute to hispid but may become glabrous; pileus, context and pore 

surface usually distinctly colored, usually brown to gray . . . . . . . . . . . . . . . . . . . . . . 5

5. Conk sessile or effused-reflexed; pileus up to 6 cm x 8 cm x 0.5 cm, villose to hirsute, 

brownish to gray in concentric zones; pore surface white becoming gray; pores usually 

daedaleoid, breaking up into teeth with age; context with a soft, dark upper layer merging into the tomentum and separated by a thin black zone from a corky, pale lower 

layer; spores 5–7 μm long . . . . . . . . . . . . . . . . . . . . . . . Cerrena unicolor (fig. 30-20)

5. Conk pileate, may have an effused lower portion; pileus hirsute to hispid or mattedstrigose to glabrous, zonate or not; pores round or elongate to daedaleoid; context 

without black zone; spores ≥8 mm long. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

6. Conk usually sessile, may be effused-reflexed, up to 3 cm thick; pileus matted-strigose 

to glabrous, buff to light brown; pores variable but usually daedaleoid; spores allantoid 

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daedaleopsis confragosa (fig. 30-18)

6. Conk usually effused-reflexed, usually ~1 cm thick or less; pileus hirsute to hispid, 

creamy-buff or brown to gray; pores round to angular, less often daedaleoid; spores 

cylindrical . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Coriolopsis
112 USDA Forest Service RMRS-GTR-335. 2016.

Coriolopsis

a. Conk sessile, effused-reflexed, or rarely resupinate, up to 6 cm x 12 cm x 2 cm; 

pileus hirsute to coarsely hispid, cream to clay-buff; pore surface white to claybuff; pores angular to daedaleoid, averaging 3 per mm but some 1 mm in diameter; 

context white to cream buff, duplex, 2–5 mm thick; spores cylindric, 9–12 μm x 

2.5–4.0 μm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Coriolopsis trogii (fig. 30-21)

a. Pileus, pore surface, and especially context darker in color; pores round to 

angular . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . b

b. Conk sessile to resupinate; pileus up to 3 cm x 6 cm x 0.3 cm but larger if fused, 

brown to gray, hirsute to tomentose; pore surface cinnamon-brown to gray, becoming darker; pores 2–4 per mm; context duplex, dense lower part up to 4 mm thick, 

light brown or golden-cinnamon and shiny where cut, temporarily black in KOH; 

spores cylindric, 8–11 μm x 2.5–4.0 μm . . . . . . . . . . . . . . . . . Coriolopsis floccosa

b. Conk sessile, but sometimes several pilei connected with common effused part; 

pileus cream to clay-buff, up to 6 cm x 12 cm x 2 cm, densely hirsute to hispid, 

yellow-brown weathering to gray, zonate or azonate; pore surface brown to gray; 

pores angular, 1–3 per mm, becoming elongated and split; context 2–10 mm thick, 

rusty to umber brown, temporarily blackish in KOH; spores cylindric, 10–16 μm x 

4–5 μm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Coriolopsis gallica

Datronia

a. Conk corky to hard, pileate to effused-reflexed; pileus glabrous; pores round to 

slightly angular, 4–5 per mm . . . . . . . . . . . . . . . . . . . . . . . . . . Datronia scutellata

a. Conk tough but flexible, effused-reflexed or resupinate; pileus velutinate to tomentose; pores angular to daedaleoid, 1–2 per mm . . . . . . . . . . . . . . . Datronia mollis

Perenniporia

a. On live buffaloberry; pores 2–3 per mm; spores 6.5–8.5 μm long 

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Perenniporia ellisiana (fig. 30-22)

a. On other hosts, pores ≥3 per mm; spores ≥9 μm long . . . . . . . . . . . . . . . . . . . . . . b

b. Typically on live ashes; conk large, up to 25 cm x 40 cm x 10 cm; pores 3–5 per 

mm; spores 9–11 μm long . . . . . . . . . . . . . . Perenniporia fraxinophila (fig. 30-3)

b. On various dead hardwoods; conks small, up to 3 cm x 3 cm x 2 cm; pores 5–7 per 

mm; spores 13–16 μm long . . . . . . . . . . . . . . . . . . . . . . . . . Perenniporia ohiensis

Trametes

a. On live or recently dead willows and sometimes other hardwoods; pileus with a 

pleasant anise odor when fresh, up to 10 cm x 16 cm x 4 cm, white, gray, or 

pale yellow-brown; context white; pore surface white to pale buff; pores round to 

angular, 2–3 per mm; spores cylindrical, 7–12 μm x 3.0–4.5 μm 

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Trametes suaveolens (fig 30-23)

a. On dead (occasionally live) willows and other hardwoods; pileus with no distinctive odor, up to 20 cm x 35 cm x 3 cm, mostly white but often drying gray or 

yellow; context cream to pale brown; pore surface white but often drying yellow 

or pale brown; pores highly variable: round, angular (1–2 per mm), daedaleoid, to 

more or less lamellate; spores cylindrical, 5–7 μm x 2–3 μm . . . Trametes elegans
USDA Forest Service RMRS-GTR-335. 2016. 113

Table 30-1—Major wood-decay fungi of the Great Plainsa

 Tree AbunFungus Name in first edition Hosts statusb dancec

Antrodia albida Trametes sepium Willows (Salix spp.) D C

Antrodia juniperina Same Junipers (Juniperus spp.), esp. L C

eastern redcedar (J. virginiana)

Antrodia malicola Trametes malicola Willows D C

Cerrena unicolor Daedalea unicolor Willows D C

Coriolopsis floccosa Trametes rigida Willows D C

Coriolopsis gallica Trametes hispida Willows D C

Coriolopsis trogii Trametes trogii Willows D C

Daedaleopsis confragosa Daedalea confragosa Willows D C

Datronia mollis Trametes mollis Willows D C

Datronia scutellata Same Hardwoods D U

Fomes fomentarius Same Birch (Betula spp.) most common D C

Fomitiporia punctata Phellinus punctatus Hardwoods, esp. green ash L+D C

(Fraxinus pennsylvanica), Siberian 

peashrub (Caragana arborescens), 

American plum (Prunus americana), 

willows, black locust (Robinia 

pseudoacacia), common buckthorn 

(Rhamnus cathartica)

Fomitiporia texana Not in first edition Junipers, esp. eastern redcedar L C

Fomitopsis cajanderi Same Conifers and hardwoods D (L) U

Fomitopsis meliae Fomitopsis melliae [sic.] Many hardwoods, including poplar L+D C

(Populus spp.), maple (Acer spp.), 

honeylocust (Gleditsia spp.), ash 

(Fraxinus spp.), occasionally conifers. 

Impt. in live peach (Prunus persica).

Fulvifomes robiniae Phellinus robineae [sic.] Black locust in central and southern L C

Great Plains

Fuscoporia gilva Phellinus gilvus Green ash, black locust, maple, willow D (L) C

Ganoderma applanatum Same Poplar, maple, honeylocust, ash L+D C

Ganoderma lobatum Same Hardwoods D U

Inonotus weirianus Phellinus weirianus Black walnut (Juglans nigra) L C

Laetiporus sulphureus Not in first edition Hardwoods L+D C

 sensu lato d

Oxyporus populinus Same Hardwoods, esp. maples L U

Perenniporia ellisiana Same Buffaloberry (Shepherdia spp.) L C

Perenniporia fraxinophila Same Ash almost exclusively L C

Perenniporia ohiensis Perenniporia ohiense [sic.] Oaks (Quercus spp.) and D (L) U

other hardwoods

Phellinopsis conchata Phellinus conchatus Hardwoods D C

Phellinus everhartii e Same Oaks L C

Phellinus igniarius sensu latof Same Birch, also ash, black walnut, L C

poplars, buckthorn, willow

Phellinus pomaceusg Phellinus tuberculosus Plum, peach, nectarine, cherry L C

(Prunus spp.)

Phellinus tremulae Same Aspen (Populus spp.) L C

Phylloporia ribis Phellinus ribis Currant and elderberry (Ribes spp.) L U

Trametes elegans Daedalea ambigua On many, common on willows D mostly C

Trametes suaveolens Same Willows, sometimes other hardwoods L C

a From (Riffle and Conway 1986; Riffle and Walla 1986; Sharon and Riffle 1986; Walla and Stack 1986).

b D = on dead trees; L = on live trees; (L) = less commonly on live trees. Note that conks on live trees may still be present after the 

tree dies. c C = common; U = uncommon.

d Laetiporus sulphureus in the broad sense. Several species have been delineated within the L. sulphureus group, and further 

collections are needed to determine which occur in the Great Plains. Laetiporus cincinnatus has been reported in Kansas. e The generic placement of Phellinus everhartii (fig. 30-5) is uncertain. It is very similar to other Phellinus species now placed in 

the Inonotus linteus group, such as I. weirianus (fig. 30-6) and I. dependens. More molecular data are needed, but available data 

suggest that it does not belong in Inonotus. Although it does not share the characters of Phellinus sensu stricto, it is retained in that 

genus here. f Phellinus igniarius in the broad sense. Many similar species have been segregated from P. igniarius over the years, but their 

distribution has not been adequately characterized. Phellinus alni is a likely species in the Great Plains because it has a wide host 

range, but it has not been specifically identified there.

g Unpublished multi-locus phylogenetic analysis shows that our Phellinus sp. on Prunus, originally named after the European 

Phellinus pomaceus, which was later synonymized with P. tuberculosus, is in fact distinct from the common P. tuberculosus in 

Europe. Furthermore, the basionym of P. tuberculosus, Boletus tuberculosus Baumgarten 1790, is illegitimate (as a later homonym 

of B. tuberculosus Bull 1783). Here the name P. pomaceus is used, although it may eventually prove to be a distinct species.
114 USDA Forest Service RMRS-GTR-335. 2016.

Figure 30-6—Conks of Inonotus weirianus on black walnut (Juglans nigra) 

(Whitney Cranshaw, Colorado State 

University, Bugwood.org., used with 

permission).

Figure 30-5— Phellinus everhartii on 

bur oak (Derek Lowstuter, North Dakota 

Forest Service, used with permission); 

(inset) close-up of a P. everhartii conk 

(James J. Worrall, U.S. Forest Service).

Figure 30-7—Oxyporus 

populinus (James A. Walla, 

Northern Tree Specialties, 

used with permission).

Figure 30-8—The sulfur shelf, 

Laetiporus sulphureus (Aaron 

D. Bergdahl, North Dakota 

Forest Service, used with 

permission).

Figure 30-9—Antrodia juniperinum on juniper in Nebraska 

(Laurie J. Stepanek, Nebraska 

Forest Service, used with 

permission).

Figure 30-10—Antrodia albida (James J. 

Worrall, U.S. Forest Service). Figure 30-11—Fomitopsis 

cajanderi (James J. Worrall, 

U.S. Forest Service). 
USDA Forest Service RMRS-GTR-335. 2016. 115

Figure 30-12—The artist’s conk, 

Ganoderma applanatum (Aaron 

D. Bergdahl, North Dakota Forest 

Service, used with permission).

Figure 30-13—Phellinus conchata

on green ash; (inset) cross section 

of decay with older conk above 

(James A. Walla, Northern Tree 

Specialties, used with permission).

Figure 30-14—Fulvifomes 

robinae; (inset) pore surface 

(James J. Worrall, U.S. Forest 

Service).

Figure 30-15—Fomitiporia 

punctata on green ash 

(James A. Walla, Northern 

Tree Specialties, used 

with permission).

Figure 30-16—Phellinus 

tremulae; (inset) pore surface 

(Aaron D. Bergdahl, North 

Dakota Forest Service, used 

with permission).

Figure 30-17—Phellinus 

pomaceus (James A. Walla, 

Northern Tree Specialties, 

used with permission).

Figure 30-18—Daedaleopsis confragosa

(James J. Worrall, U.S. Forest Service) 
116 USDA Forest Service RMRS-GTR-335. 2016.

Figure 30-19—Phellinus 

igniarius (James J. Worrall, 

U.S. Forest Service).

Figure 30-20—Cerrena unicolor (Joseph 

O’Brien, U.S. Forest Service, Bugwood.org); 

(inset) pore surface (James J. Worrall, U.S. 

Forest Service).

Figure 30-21—Coriolopsis trogii on 

willow (James A. Walla, Northern Tree 

Specialties, used with permission).

Figure 30-22—Perreniporia ellisiana on buffaloberry (James A. 

Walla, Northern Tree Specialties, 

used with permission).

Figure 30-23—Trametes spp. on plum (Aaron D. Bergdahl, North Dakota 

Forest Service, used with permission); (upper right) pore surface of Trametes

suaveolens (James J. Worrall, U.S. Forest Service); (lower right) decay 

of heartwood by Trametes spp.(Aaron D. Bergdahl, North Dakota Forest 

Service, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 117

Management

Once decay is established in a tree, little or nothing can be done to stop it. Although 

there is no cure, prevention can be effective. Avoid unnecessary wounding, such as from 

livestock, machinery, tree removal, and fire. Where pruning is needed, do it while trees 

are young and vigorous and branches are small, so that pruning wounds will close quickly. Use proper pruning techniques. Maintaining tree vigor with water during dry periods 

and fertilizer when soil testing shows a nutrient deficiency may speed wound closure and 

increase tree defenses. In general, do not apply wound dressing chemicals. There is no 

evidence that they provide any benefit, and several have been shown to be deleterious. 

Where thinning is appropriate, remove trees with large wounds, evidence of decay, and 

low vigor. If branches break, prune back to the nearest live lateral. When large wounds 

are fresh, remove any peeled bark fragments. Pruning distal to the wound may help promote wound closure.

Selected References

Bedker, P.; O’Brien, J.; Mielke, M. 2012. Rev. ed. How to prune trees. NA-FR-01-95. 

Newtown Square, PA: U.S. Department of Agriculture, Forest Service, Northeastern 

Area, State and Private Forestry. 29 p.

Gilbertson, R.L.; Ryvarden, L. 1986. Two volumes. North American polypores. Blindern, 

Norway: Fungiflora A/S. 885 p.

Lesica, P.; Atthowe, H.E.; Dugan, F.M. 2003. Incidence of Perenniporia fraxinophila 

and its effects on green ash (Fraxinus pennsylvanica) woodlands in eastern Montana, 

USA. Forest Ecology and Management. 182(1-3): 153-159.

Lowe, J.L. 1942. The Polyporaceae of New York State (except Poria). Tech. Publ. 60. 

Syracuse, NY: New York State College of Forestry. 128 p.

Lowe, J.L. 1957. Polyporaceae of North America. The genus Fomes. Tech. Publ. 80. 

Syracuse, NY: State University College of Forestry at Syracuse University. 97 p.

Overholts, L.O. 1953. The Polyporaceae of the United States, Alaska and Canada. Prepared 

for publication by J.L. Lowe. Ann Arbor, MI: University of Michigan Press. 466 p.

Riffle, J.W.; Conway, K.E. 1986. Phellinus stem decays of hardwoods. In: Riffle, J.W.; 

Peterson, G.W., eds. Diseases of trees in the Great Plains. Gen.Tech. Rep. RM-129. 

Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain 

Forest and Range Experiment Station: 83-85.

Riffle, J.W.; Ostrofsky, W.D.; James, R.L. 1981. Fomes fraxinophilus on green ash in 

Nebraska windbreaks. Plant Disease. 65(8): 667–669.

Riffle, J.W.; Sharon, E.M.; Harrell, M.O. 1984. Incidence of Fomes fraxinophilus on 

green ash in Nebraska woodlands. Plant Disease. 68(4): 322–324.

Riffle, J.W.; Walla, J.A. 1986. Perennial woodrotting fungi that cause stem decays of 

hardwoods. In: Riffle, J.W.; Peterson, G.W. eds. Diseases of trees in the Great Plains. 

Gen. Tech. Rep. RM-129. Fort Collins, CO: U.S. Department of Agriculture, Forest 

Service, Rocky Mountain Forest and Range Experiment Station: 80–82.

Rizzo, D.M.; Gieser, P.T.; Burdsall, H.H. 2003. Phellinus coronadensis: a new species 

from southern Arizona, USA. Mycologia. 95(1): 74–79.

Sharon, E.M.; Riffle, J.W. 1986. Antrodia stem decay of eastern redcedar. In: Riffle, J.W.; 

Peterson, G.W., eds. Diseases of trees in the Great Plains. Gen. Tech. Rep. RM-129. 

Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain 

Forest and Range Experiment Station: 134–135.

Vlasák, J.; Li, H.-J.; Zhou, L.-W. ; Dai, Y.-C. 2013. A further study on Inonotus linteus complex 

(Hymenochaetales, Basidiomycota) in tropical America. Phytotaxa. 124(1): 25-36.

Walla, J.A.; Riffle, J.W. 1981. Fomes fraxinophilus on green ash in North Dakota 

windbreaks. Plant Disease. 65: 669–670.

Walla, J.A. 1984. Incidence of Phellinus punctatus on living woody plants in North 

Dakota. Plant Disease. 68: 252–253.

Walla, J.A.; Stack, R.W. 1986. Stem decays of willow. In: Riffle, J.W.; Peterson, G.W., 

eds. Diseases of trees in the Great Plains. Gen. Tech. Rep. RM-129. Fort Collins, CO: 

U.S. Department of Agriculture, Forest Service, Rocky Mountain Forest and Range 

Experiment Station: 62–63. 
118 USDA Forest Service RMRS-GTR-335. 2016.

31. Thousand Cankers Disease (TCD) of Walnut

Ned Tisserat and Whitney Cranshaw

Black walnut (Juglans nigra) is native to the eastern Great Plains and has been widely 

planted throughout the region as a nut and ornamental tree. Widespread mortality of black 

walnut caused by thousand cankers disease was first noted in Colorado in 2001. The 

disease now threatens black walnuts throughout the Great Plains. 

Hosts and Distribution

The walnut twig beetle (WTB; Pityophthorus juglandis) and its fungal associate 

Geosmithia morbida cause minor to modest damage to their native host, the Arizona 

walnut (J. major). In contrast, black walnut is highly susceptible to this insect and fungus 

(known collectively as TCD), and thousands of planted black walnut in cities and rural 

locations throughout the western United States have been killed. Other native walnuts 

including butternut (J. cinerea), southern and northern California black walnuts (J. 

californica and J. hindsii, respectively), little walnut (J. microcarpa), and the introduced 

Japanese and Persian walnuts (J. ailantifolia and J. regia, respectively) are intermediate 

in susceptibility to the pathogen. 

Symptoms and Signs

Walnut trees affected by TCD initially exhibit yellowing of foliage and thinning of 

the upper crown, followed by twig and branch dieback. Progressively larger branches 

die until eventually the main stem of the tree is colonized by the beetle, and the tree is 

killed (fig. 31-1). The bark remains tightly attached to the tree, and unlike diseases caused 

by many other canker pathogens, no open-faced cankers are formed. However, external 

symptoms such as bark cracking or black sap exudation and staining of the bark surface 

may belie an underlying infection of the phloem. Sprouts may form on the trunks of dying trees, but these also succumb.

The WTB is a minute (1.5 to 2.2 mm in 

length) reddish-brown bark beetle, about 

three times as long as it is wide (fig. 31-2). 

The small entrance/emergence holes of the 

WTB may be visible on the bark of younger, 

smooth-barked branches but are not easily found on larger branches or the trunk, 

where they typically occur in bark fissures. 

The WTB rarely attacks branches smaller 

than 1 inch in diameter, so examining small 

diameter twigs is not helpful in diagnosis. The 

WTB galleries are only visible in the phloem 

after the outer bark is removed. Galleries may 

occur at various depths in the bark tissue, 

including the bark–wood interface during the 

final stages of the disease. 

The entrance holes and galleries of WTB 

are surrounded by small, diffuse dark brown 

to black cankers caused by G. morbida. These 

cankers eventually coalesce, resulting in 

necrosis of large areas of the bark (fig. 31-3). 

The light-colored (tan to yellow) fungus often 

sporulates in beetle galleries, giving them a 

dusty appearance. The barrel-shaped asexual 

spores (conidia) measure 2 μm × 5 μm and 

are formed on verticillate conidiophores 

(fig. 31-4). 

Figure 31-1—Branch dieback on black 

walnut resulting from thousand cankers 

disease (Ned Tisserat, Colorado State 

University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 119

Figure 31-2—The walnut twig beetle, 

Pityophthorus juglandis, can be identified by 

concentric rows of cuticular bumps or ridges 

(asperities) on the prothorax (Steven Valley, 

Oregon Department of Agriculture, Bugwood.

org, used with permission).

Figure 31-3—Cankers caused by Geosmithia 

morbida. The outer bark has been removed 

to expose the cankers, which form around 

walnut twig beetle galleries (Ned Tisserat, 

Colorado State University, used with 

permission).

Figure 31-4—Barrel-shaped asexual spores 

(conidia) and verticillate conidiophores of 

Geosmithia morbida (Ned Tisserat, Colorado 

State University, used with permission).

Disease Cycle

The life cycle of the WTB is not fully understood. Generations appear to overlap, and 

adults may emerge from galleries throughout 

the year during warm weather. However, 

in the Rocky Mountain region, peak adult 

emergence appears to be in late July and 

early August. The WTB carries G. morbida

on its body. The relationship of the fungus 

to the WTB is still unclear, but the fungus 

appears to begin to colonize the phloem coincident with the construction of the gallery 

beneath the bark. The infection subsequently 

causes cankers around the gallery. The total 

length of time for TCD to kill a mature walnut 

tree (from initial beetle infestation to tree 

death) is not known, but trees generally die 

within four years after symptoms develop.

Management

Currently there are no effective management strategies for TCD. Trees exhibiting 

more than 50-percent crown dieback should be removed. Wood with bark attached can 

harbor the WTB for up to 20 months and should not be moved to uninfested areas without first disinfesting by heat or another method. The effectiveness of insecticide drenches 

or injections in preventing beetle colonization is being studied. Surviving black walnuts 

in TCD-affected areas have been observed, but it is not yet known whether these trees are 

resistant. 

Selected References

Kolařík, M.; Freeland, E.; Utley, C.; Tisserat, N. 2011. Geosmithia morbida sp. nov. 

A new phytopathogenic species living in symbiosis with the walnut twig beetle 

(Pityophthorus juglandis) on Juglans in the USA. Mycologia. 103: 325–332. 

Tisserat, N.; Cranshaw, W.; Leatherman, D.; Utley, C.; Alexander, K. 2009. Black walnut 

mortality in Colorado caused by the walnut twig beetle and thousand cankers disease. 

Plant Health Progress. doi:10.1094/PHP-2009-0811-01-RS.

Utley, C.; Nguyen, T.; Roubtsova, T.V.; Coggeshall, M.; Ford, T. C.; Grauke, L.J.; 

Graves, A.D.; Leslie, C.A.; McKenna, J.; Woeste, K.; Yaghmour, M.A.; Cranshaw, W.; 

Seybold, S.J.; Bostock, R.M.; Tisserat, N. 2013. Susceptibility of walnut and hickory 

species to Geosmithia morbida. Plant Disease. 97: 601–607.
120 USDA Forest Service RMRS-GTR-335. 2016.

32. Wetwood (Slime Flux) and Alcohol Flux in 

Hardwoods

William R. Jacobi and John Ball

Bacterial wetwood is a common disease affecting the internal xylem of many softwood and hardwood trees. In some species wetwood can affect the cambium and thus 

is lethal. In other species little damage occurs. Alcohol flux is a yeast-type infection of 

surface wounds on trees that produces a foaming and sweet alcoholic smell.

Hosts and Distribution

•  Wetwood affects trees throughout the United States. It is common in elm (Ulmus

spp.), mulberry (Morus spp.), willow (Salix spp.), and aspen and cottonwood 

(Populus spp.) in the Great Plains.

•  Alcohol flux affects primarily willow, although oaks (Quercus spp.) and elms can 

also be hosts in the Great Plains

Symptoms and Signs

Wetwood can be found in the trunk, branches, and roots (figs. 32-1 through 32-4) and 

has a dark color, foul odor, high moisture content, elevated pH, decreased electrical resistance, abnormally high gas pressure, and increased levels of mobile cations. Elms, aspen, 

and cottonwoods are often infected early in their life. Dark streaks or bands may appear 

in the annual rings. Discoloration is most extensive in heartwood or older sapwood, but 

can occur in current sapwood. Foliage of affected limbs may become prematurely yellow, scorched, and wilted, although many infected trees do not exhibit these symptoms. 

Wilting may cause dieback of scattered branches, and the entire crown may decline over 

several years. Premature defoliation may occur, but causes little growth loss. “Bleeding” 

or slime-fluxing from trunk wounds, cracks, or other injuries is the most conspicuous 

symptom. Airborne bacteria, yeasts, and other fungi contaminate the sap on the outside 

of the tree, resulting in a frothy, slimy, foul-smelling liquid. Upon drying, it leaves a light 

gray to white crust (fig. 32-1). Insects may 

visit the slime but are not involved with 

the disease (fig. 32-5).

Abundant gas is produced in wetwoodaffected tissues by the fermenting action 

of bacteria on carbohydrates and other 

materials in the xylem. The gas is composed primarily of methane, nitrogen, 

carbon dioxide, and oxygen. When the gas 

is confined in the trunk, abnormally high 

pressures of up to 60 psi develop; 5 to 10 

psi is normal for healthy tissues. The accumulation of liquid under pressure results 

in a water-soaked condition that gives 

rise to the name “wetwood.” The pressure 

forces the accumulated liquid and gas out 

of the trunk through cracks in crotches, 

through pruning and other wounds, and 

through other natural openings. This liquid 

can cause localized cambial mortality and 

prevent callus and subsequent woundwood 

formation. Foliage wilts when sufficient 

quantities of the toxic liquid accumulated 

Figure 32-1—Discoloration of bark of American 

elm (U. americana) affected with wetwood 

(William R.Jacobi, Colorado State University, 

used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 121

Figure 32-2—Wetwood discoloration at branch crotch 

on American elm (William R. Jacobi, Colorado State 

University, used with permission).

Figure 32-3—Old pruning cut with wetwood slime inhibiting woundwood growth (William R. Jacobi, Colorado 

State University, used with permission).

Figure 32-4—Base of tree where wetwood slime has 

killed the grass (William R. Jacobi, Colorado State 

University, used with permission).

in the trunk wood are carried into 

the branches. Leaves first curl 

upward along their margins, then 

the petioles become flaccid, and 

finally the leaves droop and wilt.

Alcohol flux, also known as 

frothy or white flux, is not related 

to wetwood, but the symptoms 

are often confused. Alcohol flux 

is almost clear liquid although it 

is more often noticed when it has 

the appearance of foamy skim 

milk. The liquid occurs in cracks 

and fissures, usually near the base 

of the trunk flare, but can also be 

found seeping out higher on the 

trunk (figs. 32-6 and 32-7). The 

foam usually appears during the 

summer and may remain for only 

a few days or so. Alcohol flux has 

a sweet beerlike odor, whereas 

wetwood is more rancid. The sweet 

smell of alcohol flux is also attractive to insects, which may be found 

swarming around the foamy liquid. 

Alcohol flux develops just beneath 

the bark in the outermost sapwood, 

whereas wetwood is usually in the 

center of the tree.

Disease Cycle

The bacteria associated with 

wetwood disease are common soil 

and water inhabitants and probably 

enter the tree through root wounds 

early in a tree’s life. Most of the 

bacteria lie dormant and do not 

cause wetwood until the tree is 

much older. Wetwood is primarily 

associated with various facultative 

or obligate anaerobic bacteria and 

archaea; both types of organisms 

probably play a role in the production of the complex symptoms 

associated with wetwood. A related 

form of wetwood called cambial 

wetwood is more damaging to 

trees. In this situation the bacterial infestation is found in the outer xylem-cambial area 

and the fermentation causes the death of the cambium, which can cause small to large 

cankers that can girdle branches or stems (fig. 32-8).

The yeast microbes responsible for alcohol flux have not been extensively studied, 

perhaps because the disease causes minimal harm to the infected tree. Many yeast species are associated with the disease, and these organisms produce gas and ethanol during 

fermentation. 
122 USDA Forest Service RMRS-GTR-335. 2016.

Damage

Wetwood is a chronic disease 

that may contribute to general 

decline, especially of old trees 

and trees of low vigor. The 

disease affects drought-stressed 

trees most severely. It causes 

an unsightly and often foulsmelling bleeding from tree 

wounds. The disease retards or 

prevents woundwood formation 

over wounds, and therefore 

lengthens susceptibility to decay 

fungi. Dripping wetwood flux 

may kill turf beneath infected 

trees. During drought conditions 

the bacteria can move to the 

cambium and cause cankers and 

girdling of the host. These effects 

are common in recently transplanted poplars (Populus spp.) 

and related trees. 

Wetwood is responsible for 

substantial losses of wood in the 

forest products industry. Loss 

occurs through shake and frost 

cracks in living trees, checking 

and collapse in lumber and veneer during drying, and increased 

drying time in the kiln. The 

stained wood is also a serious defect in lumber and veneer graded 

on appearance. Strength properties of wetwood-infected tissues 

do not differ significantly from 

those of healthy tissues. In fact, 

wetwood tissue in living trees 

is rarely decayed and appears to 

be resistant to wood-inhabiting 

fungi.

Alcohol flux can be associated with the death of cambial 

tissue, which often turns brown 

before dying. Infected trees 

may also exhibit dieback, but 

this effect may be a result of the 

predisposing stress rather than 

alcohol flux. 

Management

Wetwood cannot be treated by chemicals. The best action is to keep trees healthy and 

not drought stressed. Drought stress intensifies the severity of disease on many hosts. 

Installation of drain tubes to lower stem pressures and remove excess liquid is not recommended because the drilling will allow the wetwood pathogens to move outward into 

healthy sapwood. The wounds also may allow entrance of decay fungi. Remove severely 

affected trees and limbs in spring. Take care to sterilize tools after each cut to prevent 

spread from diseased to healthy trees.

Figure 32-5—Wetwood or alcohol flux slime with visiting wasps (Hymenoptera: Apocrita) and bumble flower 

beetles (Euphoria inda; Whitney Cranshaw, Colorado State 

University, Bugwood.org, used with permission).

Figure 32-6—White ash (Fraxinus americana) with milky alcohol flux (James A. Walla, Northern Tree Specialties, used 

with permission).
USDA Forest Service RMRS-GTR-335. 2016. 123

Figure 32-7—Milky froth of alcohol flux 

(James A. Walla, Northern Tree Specialties, 

used with permission).

Figure 32-8—Wetwood-induced cambial canker 

on a nursery grown cottonwood (William R. Jacobi, 

Colorado State University, used with permission).

Alcohol flux infections appear to be stress related and maintaining or improving tree 

health is the only treatment. Drought and heat stress are the two most common agents 

associated with alcohol flux, so watering a tree during a summer dry period may be the 

best treatment. The symptoms often disappear following rain. The disease is also more 

common on trees that have suffered severe mechanical injury from being struck by grass 

whips and lawnmowers.

Selected References

Murdoch, C.W.; Biermann, C.J.; Campana, R.J. 1983. Pressure and composition of 

intrastem gases produced in wetwood of American elm. Plant Disease. 67: 74–76.

Murdoch, C.W.; Campana, R.J. 1981. Bacterial wetwood. In: Stipes, R.J.; Campana, 

R.J., eds. Compendium of elm diseases. St. Paul, MN: American Phytopathological 

Society: 31–34.

Murdoch, C.W.; Campana, R.J. 1983. Bacterial species associated with wetwood of elms. 

Phytopathology. 73: 1270–1273.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Toole, E.R. 1968. Wetwood in cottonwood. Plant Disease Reporter. 52: 822–823.
124 USDA Forest Service RMRS-GTR-335. 2016.

33. Witches’-Broom of Hackberry

Mark O. Harrell and Laurie J. Stepanek; revised from Mark O. Harrell 

and Frederick J. Crowe (Riffle and Peterson 1986)

Hackberry (Celtis occidentalis) is commonly planted in cities and parks in much of the 

central Great Plains. Although principally a bottomland tree, it is able to withstand a wide 

range of moisture and temperature conditions and does well in an urban environment. A 

common and often disfiguring branch disease of hackberry is witches’-broom.

Hosts and Distribution

Witches’-broom is widespread throughout the range of hackberry in the Great Plains. 

It is very common in eastern Kansas and Nebraska. The incidence and severity of witches’-brooms on individual hackberry trees can vary greatly. Sugarberry (C. laevigata) is 

rarely affected.

Symptoms and Signs

The most apparent symptom of witches’-broom is a dense clustering or “brooming” of 

twigs (figs. 33-1 and 33-2). Because brooms do not occur on branches less than one year 

old, they are not found at 

branch tips. They occasionally appear to be terminal, 

however, when a branch 

leader fails to develop fully 

or when it dies and breaks 

off. 

The first symptom in the 

development of witches’-

broom appears in the buds 

on branch portions at least 

one year old. Affected buds 

are usually smaller than 

normal, and often many 

small buds are produced 

instead of a single larger 

one. The shoots produced 

from the affected buds grow vigorously 

and produce buds that are usually more 

open and more numerous than normal.

During the next year the development 

of the broom varies depending upon the 

vigor of the affected branch. If the branch 

is vigorous and grows rapidly, its buds 

develop into branchlets similar to the original, and over a number of years a loose, 

open cluster of diseased twigs is formed. 

If the diseased branch is weak, only the 

buds at the base grow, resulting in a large 

number of abortive twigs (fig. 33-3). Over 

several years this type of development produces a compact broom with a firm woody 

center often 1 inch or more in diameter 

(fig. 33-1).

Figure 33-1—Witches’-broom of hackberry (Whitney Cranshaw, 

Colorado State University, Bugwood.org, used with permission).

Figure 33-2—Hackberry with many witches’-

brooms (Laurie J. Stepanek, Nebraska Forest 

Service, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 125

Disease Cycle

The cause of witches’-broom of hackberry is not well understood. An eriophyid mite 

Aceria celtis (syn. Eriophyes celtis) rarely found on symptomless trees is almost always 

associated with the brooming. The powdery mildew fungus Podosphaera phytoptophila

is also sometimes found. The original report of the mite-mildew association with brooms 

has been commonly misinterpreted as a cause-and-effect requirement for witches’-brooms. 

However, no experimental evidence is available to confirm or disprove the role of 

these organisms. Whether the growth abnormality results directly from mite activity, 

whether the powdery mildew is involved but is too obscure to be routinely identified, 

or whether a virus-like entity is responsible remains uncertain. 

Damage

Witches’-brooms on hackberry are more unsightly than harmful to the tree. Brooms 

can cause branches to break more easily, and extensive brooming can reduce the vigor of 

the tree, but trees are seldom seriously damaged.

Management

No effective control measures are known for this disease. Pruning back affected twigs 

to sound wood can improve the tree’s appearance.

Selected References

Kellerman, W.A.; Swingle, W.T. 1888. Branch knot of the hackberry. Annual Report of 

the Kansas Experiment Station. Manhattan, KS: Kansas State Agricultural College: 

302–315.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Snetsinger, R.; Himelick, E.B. 1957. Observations on witches’-broom of hackberry. Plant 

Disease Reporter. 41: 541–544.

Figure 33-3—Witches’-broom with new and abortive twigs (William M. Ciesla, Forest Health 

Management International, Bugwood.org).
126 USDA Forest Service RMRS-GTR-335. 2016.

34. Dutch Elm Disease 

Kelsey L. Dunnell and Aaron D. Bergdahl

Dutch elm disease (DED) is a systemic vascular wilt disease of elm (Ulmus spp.) 

caused by two closely related fungi, Ophiostoma ulmi (Buisman) Nannf. and O. novoulmi Brasier, which are spread by bark beetles and through root grafts. The fungi have 

been spreading across North America since the 1920s and have killed most of the native 

elm trees in the United States over the past century.

Hosts and Distribution

Ophiostoma ulmi and O. novo-ulmi are exclusively pathogens of trees in the elm 

family (Ulmaceae). Elm species native to the Great Plains include: American elm (U. 

americana), rock elm (U. thomasii), slippery (red) elm (U. rubra), winged elm or wahoo 

(U. alata), and cedar elm (U. crassifolia). The first four species are very susceptible to 

the DED fungi and are quickly killed; however, the cedar elm, a native to the southern 

Great Plains, has some resistant individuals. Several exotic elms have been planted in 

the Great Plains. The most common is Siberian elm (U. pumila), which has been widely 

used in shelterbelts, on farmsteads, and in urban settings, and is considered moderately 

tolerant to DED. Among other Asiatic elms, U. davidiana var. japonica, U. laciniata, and 

U. villosa contain a high proportion of resistant individuals; the Chinese or evergreen elm 

(U. parvifolia) is also considered highly resistant to DED. A range of disease resistance 

occurs within many elm species. Tree breeding programs in the United States, Canada, 

and Europe continue to work on developing selections and elm hybrids that are resistant 

to DED.

Symptoms and Signs

New infections established by bark beetles (fig. 34-1) on DED-susceptible elm species, such as American elm, usually involve symptom development on a single branch. 

The first symptom of DED is yellowing of foliage, followed by wilting, browning, and 

early defoliation. These symptoms progress downward from the point of infection to all 

lower branches and eventually the main trunk and root system; the entire tree finally wilts 

and dies (fig. 34-2). This progression of symptoms may develop in a single season in 

highly susceptible species, but often takes longer (two or more years). However, observations indicate that when trees are infected via root grafts, wilting often begins in spring in 

lower branches and progresses upward, leading to mortality during that year; some trees 

may persist to the next year. 

The fungi are strictly vascular parasites, invading the xylem vessels of the host tree 

and causing disruption of water movement in the tree, which leads to brown streaking in the wood just under the bark (fig. 34-3). Other wilt-causing pathogens may 

also cause streaking of the wood, so this symptom is not entirely diagnostic for DED. 

Figure 34-1—Three species of bark beetle found in the Great Plains that spread DED: (A) Native 

elm bark beetle, Hylurgopinus rufipes; (B) Smaller European elm bark beetle, Scolytus multistriatus 

(black arrow indicates prominent abdominal spine); (C) Banded elm bark beetle, Scolytus schevyrewi, 

(black arrow indicates light banding pattern on elytra) (Guy Hanley, Minot State University, used with 

permission).
USDA Forest Service RMRS-GTR-335. 2016. 127

Figure 34-2—Infected elm trees showing dying back of 

branches with yellowing, wilting, and browning leaves 

(Aaron D. Bergdahl, North Dakota State University, used 

with permission).

Figure 34-3—A healthy and infected 

branch of American elm. Brown streaking 

in the wood (xylem) indicates presence of 

the fungus in host vascular tissue just under the bark (Jared M. LeBoldus, Oregon 

State University, used with permission).

Figure 34-4—Dark-stemmed synnemata supporting a pale globose ball of sticky liquid containing the 

spores of Ophiostoma ulmi or O. novo-ulmi (Kelsey 

L. Dunnell, North Dakota State University, used with 

permission).

Positive diagnosis requires a laboratory test in which wood chips from fresh symptomatic branches are placed on solid culture media for growth. Within seven to 10 days 

dark fungal stalks (synnemata; 2 to 5 mm high) grow on the wood chips and support a 

pale, globose ball of sticky liquid containing asexual spores of O. ulmi or O. novo-ulmi 

(fig. 34-4).

Disease Cycle

In North America, there are three known vectors for the DED fungus: Hylurgopinus 

rufipes (native elm bark beetle), Scolytus multistriatus (smaller European elm bark 

beetle), and, most recently confirmed, Scolytus schevyrewi (banded elm bark beetle) 

(fig. 34-1). The cycle of infection by 

the causal fungus is closely linked to 

the life cycles of the insect vectors. 

These DED fungi are uniquely adapted 

for, and dependent on, overland 

transmission by elm bark beetles. 

The beetles breed under the bark 

of weakened or recently killed elm 

branches, whole trees, or cut logs. If 

the DED fungus is present at breeding 

sites, newly emerged beetles will carry 

spores of the pathogen to healthy elms 

and introduce the disease when feeding on the xylem of twigs and small 

branches in the crown. The beetles can 

fly up to 0.25 miles in search of feeding or breeding sites, but they may be 

blown many miles by winds.
128 USDA Forest Service RMRS-GTR-335. 2016.

Damage

In the Great Plains, the American elm grows naturally in riparian areas. Because of 

its many desirable qualities, it was also frequently planted as ornamental and boulevard 

trees on farmsteads and in communities as well as in windbreaks. Today, most of the 

American elms in the Great Plains have been lost to DED. However, substantial numbers 

still remain in the very northern and southern parts of the region, as well as in communities that practice strict sanitation methods to manage the disease. This strict sanitation 

effort does not come without cost, but it allows communities to maintain thriving elm 

populations and also minimizes tree removal and replanting costs. The intangible costs 

of losing a well-adapted tree species for farmstead and windbreak plantings and the loss 

of a dominant species in riparian forest ecosystems are difficult to quantify, but certainly 

highly significant.

Management

In urban areas, a strict sanitation program can be extremely effective in slowing the 

spread of DED. Sanitation consists of removing sources of beetle habitat, which includes 

pruning and removing any dead or damaged branches and disposal of infected elm wood 

by burning, burying, chipping, or debarking. To limit the spread through root grafts to adjacent elm trees, the area around infected elm trees should be trenched (midway between 

trees) to a depth of 36 to 40 inches, and the trench should completely encircle infected 

trees. For these sanitation efforts to be effective, it is important that newly infected 

trees be identified quickly, removed, and destroyed promptly. Some cities have strict 

ordinances requiring prompt removal of infected trees and enforce bans on elm firewood 

with intact bark. The reward for these DED management efforts is a well-maintained elm 

population and a rate of tree loss reduced to a level the community can afford, allowing 

for gradual replacement with a diversity of species adapted to the region. 

Unfortunately, in rural areas, windbreaks, and native woodlands, little can be done to 

forestall the loss of elms susceptible to DED. However, individual high-value trees may 

be protected by injection of fungicides at one- to three-year intervals into the tree root 

flares for the entire lifespan of the tree. The level of protection provided by fungicide 

treatments depends on tree uptake. Although treatment can be relatively expensive and 

repeated wounding from injections may seriously damage the tree, the value of a mature 

boulevard, landscape, or specimen tree may justify the cost. Some fungicide injections 

are labeled for therapeutic treatment, but the success of treating after infection is poor; 

treatment may nonetheless be worth trying on very high-value trees. 

Although no elm species is totally resistant to the DED fungi, some of the Asiatic 

elms, as well as selections of American and European elms, have high levels of tolerance 

or resistance to infection by O. ulmi and O. novo-ulmi. Today, elm selections and hybrids 

from disease-resistant parents have been propagated and recently released for the commercial trade. Most of the new available cultivars are DED resistant. It is important to use 

cultivars best suited to the local environmental conditions in which they will be planted. 

Contact your local extension agent or arborist for current information and availability of 

DED-resistant cultivars.

Selected References

D’Arcy, C.J. 2000. Dutch elm disease. 2005 update. The Plant Health Instructor.

doi:10.1094/PHI-I-2000-0721-02. 

Gibbs, J.N. 1978. Intercontinental epidemiology of Dutch elm disease. Annual Review of 

Phytopathology. 16: 287–307.

Kondo, E.S.; Hiratsuka, Y.; Denyer, W.B.G. 1982. Proceedings of the Dutch elm disease 

workshop and symposium. Winnipeg, MB: Manitoba Department Natural Resources. 

517 p.

LeBoldus, J.M.; Bergdahl, A.; Knodel, J.; Zeleznik, J. 2013. Dutch elm disease in North 

Dakota: a new look. Fargo, ND: North Dakota State University Extension Service. 4 p.

Sinclair, W.A.; Campana, R.J., eds. 1978. Dutch elm disease: perspectives after 60 years. 

Ithaca, NY: Cornell University Agricultural Experiment Station. Search (Agriculture) 

8(5): 1–52.
USDA Forest Service RMRS-GTR-335. 2016. 129

35. Oak Wilt 

Jennifer Juzwik and David N. Appel 

Oak wilt, caused by the fungus Ceratocystis fagacearum, is known to exist only in 

the central and eastern United States, although susceptible hosts and apparently suitable 

climate conditions exist in other parts of the country. Oak wilt was likely killing trees 

in Wisconsin and Minnesota as early as the 1890s based on historical accounts, but the 

causal fungus was not discovered and described until 1944 in Wisconsin. The origin of 

the fungus is unknown, although more recent evidence suggests that it may be an introduced pathogen from Central America, South America, or Mexico.

Hosts and Distribution

The disease is currently found in an area bounded by Minnesota to Pennsylvania in 

the north and Texas to South Carolina in the south. A recent new find (2008) in upstate 

New York was quickly treated and is now considered eradicated. Within the Great 

Plains states, the disease is widespread and common in Texas but is reported in eastern 

Oklahoma, Kansas, Nebraska, and South Dakota. 

More than 33 species of oaks (Quercus spp.) as well as six other close relatives are 

known to be susceptible to C. fagacearum based on artificial inoculation studies or 

documented natural infections. Species of the red oak group, such as black (Q. velutina), 

blackjack (Q. marilandica), northern red (Q. rubra), pin (Q. palustris), and Shumard 

(Q. shumardii), are considered to be highly susceptible to the pathogen compared to 

members of the white oak group, such as bur (Q. macrocarpa), post (Q. stellata), dwarf 

chinkapin (Q. prinoides), and white oak (Q. alba), which display moderate to high levels 

of resistance. A range of susceptibility occurs in semi-deciduous live oaks (Quercus 

fusiformis and Q. virginiana) in Texas, depending on timing of infection, weather conditions, and host genetics. 

Symptoms and Signs

Foliar symptoms of oak wilt occur any time from late spring to late summer or early 

fall. Symptom expression and progression are different in red oaks, white oaks, and live 

oaks.

Early foliage symptoms in red oaks are characterized by a subtle, off-green to bronze 

color. Necrosis on wilting leaves starts at the leaf margins and tip, progressing inward to 

the main vein and the base of the leaf. Foliage symptoms commonly occur first in the upper portion of the tree crown (fig. 35-1). Progression of the disease throughout the crown 

can be rapid, resulting in complete tree wilt within several weeks to several months. Red 

oaks infected later in the growing season may produce new shoots in the lower crown the 

next spring, but these will quickly die. Infected red oaks do not recover from the disease. 

Light bluish-gray to darker streaking develops in the outermost layer of wood of branches 

with recently wilted leaves or in the main stem of a completely wilted tree, although such 

streaking is often hard to detect (fig. 35-4A). Groups of red oaks may die over several 

years if grafted roots occur between closely spaced trees.

Foliar symptoms in moderately resistant white oak trees occur on scattered branches in 

the tree crown (fig. 35-2). Discoloration on wilting white oak leaves is more variable than 

that on red oaks. Reddish-brown to brown discoloration on bur and white oak starts at 

the leaf margin and toward the tip, but may often be limited to one side of the main vein. 

Symptomatic leaves of bur oak may be confused with those of bur oak blight, which is 

caused by the fungus Tubakia iowensis (see chapter 3). The veinal chlorosis and necrosis 

of live oak leaves are unique and diagnostic for the disease (fig. 35-3). Symptoms in 

white oak species develop more slowly than in red oaks, but time to tree death differs by 

species. Diseased live oaks in Texas usually die within three to eight months of infection. 

Moderately resistant oaks, such as bur oak, may die within two to four years of infection. 

In the highly resistant white oak, C. fagacearum infections cause dieback in one or more 

branches, but affected trees may not die for decades if at all. Such trees may be misdiagnosed as having symptoms of oak decline. Dark brown to blackish streaks in the outer 
130 USDA Forest Service RMRS-GTR-335. 2016.

xylem are visible when the 

bark is removed from an 

oak wilt-infected branch 

of some white oak species. 

This symptom appears as a 

ring of discolored tissue in 

the outer xylem in a cross 

section of the same branch 

(fig. 35-4B).

Foliar symptoms in 

live oak are unique and 

diagnostic for the disease. 

The veins turn chlorotic and 

eventually necrotic while 

the remainder of the leaf remains green. Diseased live 

oaks in Texas usually die 

within three to eight months 

of infection (fig. 35-3).

Mycelial masses, or fungal mats, of the fungus may appear on the inner bark and outer 

wood within several months or the spring after a tree dies from oak wilt (fig. 35-5). These 

occur most commonly in red oaks and occasionally, seldom, or never on species in the 

white oak group. Oak wilt mats are ephemeral in nature and can be difficult to detect on 

trees with deeply furrowed bark. A vertical crack occurs when special pad-like structures 

on the light to dark gray-colored mats rupture the bark. The ripe-fruit odor produced by 

the fungus is attractive to insects, particularly to sap beetles (Coleoptera: Nitidulidae).

Disease Cycle

The oak wilt fungus is spread from diseased oaks to healthy ones through aboveground 

transmission by insect vectors or through connected root systems. Aboveground spread to 

a healthy tree occurs when sap beetle vectors carrying spores of the oak wilt fungus visit 

xylem-penetrating wounds, generally within three days of the wounds’ creation. Such 

spread results in new oak wilt centers (or “pockets”). Susceptible wounds can be created 

Figure 35-1—Foliar and crown symptoms of oak 

wilt in northern red oak (Jennifer Juzwik, U.S. 

Forest Service).

Figure 35-2—Foliar and crown symptoms of 

oak wilt in bur oak (Jennifer Juzwik, U.S. Forest 

Service).

Figure 35-3—Foliar and crown symptoms of oak wilt in Texas 

live oak (David N. Appel, Texas A&M University, used with 

permission).
USDA Forest Service RMRS-GTR-335. 2016. 131

by construction equipment, wind damage during storms, pruning or tree felling activities, 

vandalism, or any agent that penetrates through the bark and into the sapwood. 

Fresh wound volatiles attract the principal sap beetle vectors (Carpophilus sayi and 

other Coleopterous spp.). Once the fungus has entered the vascular system of the tree, its 

asexual spores (conidia) are spread internally by sap flow to distant portions of the tree. 

Oaks respond to infection by producing balloon-like structures (tyloses) that plug the 

xylem vessels. This response, along with additional plugging of the vessels by metabolic 

fungal products, disrupts the sap flow, causing symptoms in the diseased tree. After the 

tree wilts, the fungus grows outward in its hyphal form from the xylem vessels to the 

bark-wood interface (the cambium), where the fungal mass accumulates to form the oak 

wilt mats. 

Sexual spores (ascospores) are produced on mats when sap beetles introduce spores 

of the opposite mating type of the fungus. Sap beetles pick up spores of the fungus while 

feeding on or crawling across the sporulating mats. As the mats age, the beetles emerge, 

carrying the fungus inoculum to infect wounds on healthy trees.

Shared root systems of clonally propagated live oak species allow for belowground 

spread of C. fagacearum in those species. Naturally occurring root grafts that connect the 

vascular system of neighboring oak tree roots also provide a pathway for fungus spread 

Figure 35-4—Vascular streaking in outer sapwood of northern red oak: (A) Tangential view of bluegray streaking; (B) Brown discoloration in outer growth ring of stem cross section (Jennifer Juzwik, 

U.S. Forest Service).

A B

Figure 35-5—Oak wilt fungal mat (dark gray) on the bark–wood interface of 

northern red oak with pressure pad in the center (Jennifer Juzwik, U.S. Forest 

Service).
132 USDA Forest Service RMRS-GTR-335. 2016.

among oak species. Inter-tree grafts most commonly form between oaks of the same species; however, grafts can also form between different oak species. Root grafts occur most 

frequently in light textured and deep soils in relatively flat terrain. Root sprouting occurs 

in live oaks and results in new clonal trees (ramets) that share a common root system. 

In the roots of an infected tree, conidia may be translocated through root grafts to an 

adjacent healthy oak or through the shared root system of clonally propagated live oak 

species, resulting in belowground spread. Upward movement of the fungus from the roots 

to the aboveground portion of the tree through the vascular system leads to the wilting 

process previously described. The fungus can continue to spread to nearby oaks through 

connected roots, leading to expansion of the disease center. 

Damage

Once infected, red oak species and many white oak species die from oak wilt. 

Although an infected, highly resistant white oak may not die from oak wilt, the death of 

major limbs may render the tree undesirable as a landscape or specimen tree. Oak wilt is 

a devastating disease in portions of the oak wilt range where epidemics are ongoing. Tens 

of thousands of oaks die annually from oak wilt in the Upper Midwest and millions of 

oaks have died in Texas since discovery of the disease there. In contrast, oak wilt occurs 

sporadically in the Appalachian Mountains and the Ozark Mountains. The small, isolated 

disease centers that result from overland spread often die out because of the diversity of 

species, topography, or other site limitations. Regardless, the disease can cause significant losses in urban and community forests in these regions as well as other parts of the 

disease range. When consistent, integrated, and sustained suppression programs are used, 

oak wilt has proven to be a manageable disease in states experiencing ongoing epidemics.

Management

 Management of oak wilt is based on general approaches that either prevent 

aboveground spread or stop belowground spread of the fungus, whether in a residential 

lot or a rural forest. The range of oak wilt in the United States cuts across many forest 

types and dominant Quercus species, so that practices or tools to achieve consistent 

management are not appropriate for every circumstance owing to varying site conditions, 

costs, and predicted environmental disruption. Availability of equipment may also be a 

factor in tool selection. In general, effective oak wilt management is a result of using a 

variety of tools or strategies that limit the spread of the causal fungus.

Early diagnosis of oak wilt is critical to managing the disease effectively and to justify 

the costs of treatment. Thus, annual monitoring for the disease is important in localities 

known to have oak wilt. When infected oaks are found, approaches can be developed 

based on resources available, landowner concern, and established priorities for management. When possible, remove recently wilted red oaks to eliminate sources of inoculum 

for insect vectors. Removal and proper disposal should occur before mats are produced in 

the late winter or spring of the year following tree wilt. Logs from diseased oaks should 

not be removed from the site without being properly treated. Long-distance transmission 

of the fungus by insects emerging from transported logs has resulted in major extensions 

of the oak wilt range.

Prevent infection of healthy oaks by avoiding wounding of trees, particularly in spring 

and early summer, when risk of aboveground spread is highest. Community ordinances 

may require use of protective measures during construction activities. Utility and arboriculture companies may restrict line-clearing and tree trimming during critical time 

periods and apply wound treatments when cutting is unavoidable. Educational outreach, 

such as billboards and public service announcements, raise public awareness of the 

need to prevent wounding and the hazards of moving oak firewood and logs. Municipal 

ordinances that mandate removal and treatment of dead oaks in oak wilt areas are also 

helpful.
USDA Forest Service RMRS-GTR-335. 2016. 133

Belowground spread of the oak wilt fungus accounts for the vast majority of oaks that 

die annually from the disease. Common root systems or grafted roots can be disrupted by 

using mechanical or chemical means to stop this spread. Cutting roots with a trencher to 

a depth of at least 48 inches or a vibratory plow with a 60-inch-long blade is an effective 

method in areas where soils and sites are suitable for the equipment. Shallow rocky soils 

and even-layered rock in Texas have necessitated the use of rock saws able to obtain the 

same depths for cutting oak roots. Chemical fumigants injected into 18-inch-deep holes 

evenly spaced along a line have been used in the past to create a narrow strip of dead 

roots between oak wilt-affected and healthy trees. Chemical toxicity and practical application concerns have greatly reduced or eliminated use of fumigants by practitioners. 

Most failures of root cutting or disruption treatments to control oak wilt are attributed 

to insufficient depth of treatment or poor placement of the treatment line. Several models 

have been developed to guide management foresters in placing the treatment line. The 

general approach is to place the line far enough away from the diseased trees that there 

is low probability of the fungus being in the roots of the symptom-free trees outside the 

treatment line. However, the line should be close enough that the fungus will die before 

reaching the treatment line. It is assumed that the fungus will die in cut or otherwise 

killed roots within several years; then re-grafting can occur. There is some evidence 

that inserts in the trench may improve their performance by acting as a barrier to regrafting of roots, but such materials have had only limited use. Felling of all oaks within 

a treatment line and even extraction of the stumps has been used to increase success of 

treatments in some state programs. 

Systemic fungicides injected into the vascular system of symptom-free, high-value 

oaks in close proximity to diseased trees have been used to suppress disease development 

in trees at high risk of infection, but will not limit belowground spread of the pathogen. 

There are several methods available to inject trees, but only approved, labeled fungicides 

should be used to control oak wilt. Also, use an injection method only if it has been 

proven effective through documented scientific study, is relatively easy to use, and is safe 

for the applicator. Injection and subsequent pruning of symptomatic (<30 percent crown 

wilt) live, bur, and white oak can also suppress disease development, maintain aesthetic 

appeal, and extend longevity of high-value trees.

The management of oak wilt requires a sustained, long-term commitment in order to 

ultimately contain the pathogen and save trees. In this regard, a management plan may 

also include replanting. If so, any non-oak tree species can be used as long as they are 

well-suited for the site. If oaks are preferred, then consider only members of the white 

oak group.

Selected References

Billings, R.F.; Appel, D.N., eds. 2009. Proceedings of the National Oak Wilt Symposium; 

2007 June 4–7; Austin, TX. Publ. 166. College Station, TX: Texas Forest Service. 

267 p.

Juzwik, J.; Appel, D.N.; MacDonald, W.L.; Burks, S. 2011. Challenges and successes in 

managing oak wilt in the United States. Plant Disease. 95: 888–900.

O’Brien, J.G.; Mielke, M.E.; Starkey, D.; Juzwik, J. 2011. How to identify, prevent, and 

control oak wilt. NA-FR-01-11. St. Paul, MN: U.S. Department of Agriculture, Forest 

Service, Northeastern Area State and Private Forestry. 30 p. http://na.fs.fed.us/pubs/

howtos/ht_oakwilt/identify_prevent_and_control_oak_wilt_print.pdf. (May 14, 2014).

Pokorny, J. 1999. How to collect field samples and identify the oak wilt fungus in the 

laboratory. NA-FR-01-99. St. Paul, MN: U.S. Department of Agriculture, Forest 

Service, Northeastern Area State and Private Forestry. 12 p. http://www.na.fs.fed.us/

spfo/pubs/howtos/ht_oaklab/toc.htm. (May 14, 2014).
134 USDA Forest Service RMRS-GTR-335. 2016.

36. Verticillium Wilt

Mark O. Harrell and Laurie J. Stepanek; revised from David S. 

Wysong and Mark O. Harrell (Riffle and Peterson 1986)

Two closely related species of fungi, Verticillium albo-atrum and V. dahliae, are the 

cause of Verticillium wilt in hundreds of woody and herbaceous plant species around the 

world. Both species can infect and kill trees and shrubs, but wilt caused by V. dahliae is 

more common. 

Hosts and Distribution

Verticillium wilt affects more than 300 species of plants, including food and fiber 

crops, annual and perennial ornamentals, and landscape trees. The disease occurs in forest 

stands but is far more destructive in landscape plantings. Valuable ornamental trees can 

be killed or severely damaged. Verticillium wilt occurs in every country in the world and 

in every state in the United States. Susceptible trees include apple and crabapple (Malus

spp.), ash (Fraxinus spp.), catalpa (Catalpa spp.), cherry (Prunus spp.), elm (Ulmus

spp.), linden (Tilia spp.), black locust (Robinia pseudoacacia), Kentucky coffeetree 

(Gymnocladus dioicus), magnolia (Magnolia spp.), maple (Acer spp.), pin oak (Quercus 

palustris), red oak (Q. rubra), poplar (Populus spp.), redbud (Cercis spp.), Russian-olive 

(Elaeagnus angustifolia), serviceberry (Amelanchier spp.), and smoke tree (Cotinus spp.). 

Some species and cultivars of apple, crabapple, linden, poplar, and serviceberry may be 

resistant depending on the strain of Verticillium in the soil. 

Symptoms and Signs

Verticillium wilt in trees has acute 

and chronic symptoms. Acute symptoms 

include leaf wilting, curling, and drying 

on individual branches (figs. 36-1, 36-2 

and 36-3). In maple, catalpa, and elm 

these symptoms are sometimes preceded 

by a general yellowing or reddening of 

the foliage. Trees may begin to show 

leaf symptoms as early as March or as 

late as November. If early symptoms on 

a few branches go unnoticed, sudden 

wilting of the entire crown may be the 

first symptom seen. Chronic symptoms 

include a slowing of twig growth, sparse 

foliage, stunted leaves, abnormally 

heavy seed crops, and branch dieback. 

Some trees such as maple may have 

long areas of dead bark on branches 

and the trunk. Other trees, such as elms, 

are more tolerant to infection and may 

decline slowly over several years.

Vascular streaking is another symptom of Verticillium-infected trees. In 

branches with advanced stages of wilt, 

the sapwood discolors in the form of 

bands or streaks that follow the grain of 

the wood (fig. 36-4). Discoloration occurs most frequently in the earlywood of 

the current season’s growth. In trees that 

wilt in early summer, the discoloration 

may not be noticeable when the branch 

is examined in cross section. However, 

Figure 36-1—Wilting branch on catalpa caused 

by Verticillium wilt (John Ball, South Dakota State 

University, used with permission).

Figure 36-2—Norway maple leaf (Acer platanoides) 

with Verticillium wilt symptoms (Mark O. Harrell, 

Nebraska Forest Service, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 135

it is usually seen as fine streaks on the 

surface of the sapwood when the bark is 

peeled carefully from a wilted branch. In 

branch cross sections, the discoloration appears as a series of dots in a wood ring. In 

some cases, the dots are so abundant that 

the entire ring appears discolored. Trees 

that have been infected for more than a 

year may have discoloration in more than 

one growth ring (fig. 36-5).

In severely wilted trees the discoloration in the sapwood may be abundant and 

extend to the tips of wilted branches. In 

others the discoloration may be limited to 

the trunk sapwood, or it may extend only 

a few inches into the basal portions of 

wilted branches. In maple, the discolored 

wood is light to dark green. In catalpa it is 

purplish pink, changing to bluish brown 

upon drying. In elms and many other tree 

species, it is light to dark brown. Ash 

wood often shows no streaking, or the 

streaking may be very light, even when 

severely infected.

Disease Cycle

Verticillium is a soil-borne fungus that 

invades trees through roots. When roots in 

infested soil grow close to Verticillium hyphae or resting structures (microsclerotia), 

the hyphae in the soil or from germinating 

microsclerotia penetrate intact nonwoody 

roots or enter both woody and nonwoody 

roots through wounds. After entering 

roots, the fungus spreads through the tree 

either by growth of hyphae or by spores 

(conidia) transported by upward movement of xylem sap. Verticillium can exist 

for several years in roots and trunks of 

trees killed by the disease. 

Figure 36-3—Norway maple with branches killed 

by Verticillium wilt (Mark O. Harrell, Nebraska 

Forest Service, used with permission).

Figure 36-5—Norway maple with discolored 

sapwood and loose bark from Verticillium wilt 

(Mark O. Harrell, Nebraska Forest Service, used 

with permission).

Figure 36-4—Verticillium wilt streaking in Norway maple (Mark O. Harrell, Nebraska Forest 

Service, used with permission).
136 USDA Forest Service RMRS-GTR-335. 2016.

Microsclerotia allow the fungus to persist in soil for many years away from a parasitized host. These resting structures are most abundant in the top 12 inches of soil. Conidia, 

which are important in fungal movement within the tree, do not survive in soil for more 

than a few weeks.

The movement of Verticillium to new areas occurs mostly by the movement of microsclerotia. They can be carried in root balls, on bare roots of infected trees, and in wood 

chips from trees killed by the disease. The microsclerotia contaminate the soil where the 

trees are planted or wood chips are placed. Because microsclerotia can exist for many 

years in infected wood, trees killed by Verticillium should not be used as mulch. Within 

nursery fields, microsclerotia can be spread by normal tillage operations and in soil that 

adheres to equipment that has been used in an infested field. 

Damage

Damage caused by Verticillium wilt is variable and depends upon age and species of 

the host affected. Trees with trunk diameters of 1 to 2 inches can be killed within one 

year of infection. Older trees may live many years after infection, but they normally decline over time. The disease typically has a greater impact on nursery seedlings and trees 

in landscape plantings compared to forest stands.

Management

Prevention is the best method for controlling Verticillium wilt. Avoid planting 

susceptible trees in areas where other plants are known to have died from the disease. 

Trees resistant or immune to Verticillium include arborvitae (Thuja occidentalis), 

beech (Fagus spp.), birch (Betula spp.), butternut (Juglans cinerea), fir (Abies spp.), 

ginkgo (Ginko biloba), hackberry (Celtis spp.), hawthorn (Crataegus spp.), hickory 

(Carya spp.), honeylocust (Gleditsia triacanthos), hophornbeam (Ostrya virginiana), 

juniper (Juniperus spp.), larch (Larix spp.), linden, mountain-ash (Sorbus spp.), 

mulberry (Morus spp.), bur oak (Q. macrocarpa), white oak (Q. alba), pear (Pyrus

spp.), pine (Pinus spp.), spruce (Picea spp.), sycamore (Platanus occidentalis), walnut 

(Juglans spp.), and willow (Salix spp.). Some species and cultivars of apple, crabapple, 

linden, poplar, and serviceberry are resistant. 

Trees with recent or mild symptoms of Verticillium wilt should not be removed immediately because they may recover to some degree after cultural treatments that improve 

environmental conditions and strengthen tree vigor. Water stress increases a tree’s 

susceptibility to Verticillium wilt. Watering appropriately during periods of drought and 

using a mulch bed of wood or bark chips to improve soil conditions can reduce water 

stress and increase resistance to the disease. Do not use mulch from trees that were killed 

by Verticillium, because microsclerotia can exist for many years in the infected wood. 

Branches with severe symptoms should be pruned out, but those with mild symptoms 

could be left to see if they recover. Fungicide treatments are not effective. 

Selected References

Ash, C.L. 2001. Verticillium wilt of trees and shrubs. In: Ash, C.L., ed. Shade tree wilt 

diseases. Proceedings, Wilt diseases of shade trees: a national conference. St. Paul, 

MN: APS Press: 157–163.

Carter, J.C. 1979. Diseases of Midwest trees. Urbana, IL: University of Illinois Press. 

168 p.

Himelick, E.B.; Neely, D. 1981. Verticillium wilt. In: Stipes, R.J.; Campana, R.J., eds. 

Compendium of elm diseases. St. Paul, MN: American Phytopathological Society: 22–24. 

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: Cornell 

University Press. 660 p.

Smith, L.D. 1979. Verticillium wilt of landscape trees. Journal of Arboriculture. 5(9): 

193–197.
USDA Forest Service RMRS-GTR-335. 2016. 137

37. Yellows Disease of Ash, Lilac, and Elm

James A. Walla and Les Koch; partially revised from 

Wayne A. Sinclair and David S. Wysong (Riffle and Peterson 1986)

Yellows diseases are caused by phytoplasmas (bacteria lacking cell walls). 

Phytoplasmas occur systemically in phloem tissue of their host plants and in their 

insect vectors (primarily leafhoppers and planthoppers). Ash yellows (AshY) and lilac 

witches’-broom (LWB) are caused by ‘Candidatus Phytoplasma fraxini’. Elm yellows 

(EY, previously known as elm phloem necrosis) is caused primarily by ‘Ca. P. ulmi’, but 

it was found to be caused by a member of the clover proliferation phytoplasma group in 

one area in northern Illinois. There is a wide range of aggressiveness among isolates of 

many phytoplasmas. Such has been demonstrated for AshY phytoplasmas and would be 

expected for LWB and EY.

Hosts and Distribution

AshY phytoplasma hosts include at least 13 ash (Fraxinus) species, including black 

(F. nigra), green (F. pennsylvanica), white (F. americana), and velvet ash (F. velutina), 

and 20 lilac (Syringa) species, including common (S. vulgaris), dwarf Korean (S. meyeri), 

Japanese tree (S. reticulata), late (S. villosa), and Manchurian lilac (S. pubescens subsp.

patula). Naturally-infected EY phytoplasma hosts include most native North American 

elm (Ulmus) species—American (U. americana), cedar (U. crassifolia), September 

(U. serotina), slippery (red) (U. rubra), and winged elm (U. alata)—Chinese elm (U. parvifolia), and the American-Asian hybrid slippery × Siberian elm (U. pumila). In New 

York, the white-banded leafhopper (Scaphoideus luteolus) is a known vector of the EY 

phytoplasma. In Pennsylvania, the spittlebug Lepyronia quadrangularis and the leafhopper Latalus spp. have been shown to be vectors. Leafhoppers in the genus Scaphoideus

are probably the principal vector of the AshY phytoplasma.

AshY occurs primarily in North America. In the Great Plains, it is present in most sites 

with mature ash from Alberta to Colorado and Manitoba to Oklahoma. LWB is known in 

eastern and central United States. In the Great Plains, it is known only from North Dakota 

and Oklahoma. EY occurs in eastern and central United States and was present in southern Ontario; it also occurs in Europe. It is known from the eastern Great Plains states and 

Wyoming, but is rarely recognized. States and provinces in the Great Plains where these 

diseases are not known have not been adequately sampled.

Symptoms

Symptoms of AshY and LWB may include any combination of reduced growth, 

deliquescent branching, progressive branch death, abnormal roots, chlorotic or deformed 

leaves, and witches’-brooms (fig. 37-1). Witches’-brooms are the only diagnostic 

symptom of AshY and LWB. Witches’-brooms have fine, short, and abnormally upright 

branches; branches growing from leaf axils; and dwarfed, deformed leaflets. Witches’-

brooms are rare on infected green ash in the Great Plains, but are common on trunks of 

trees that have top dieback or have been cut. In lilac, witches’-brooms are common on the 

most susceptible hosts (fig. 37-2). Witches’-brooms are rare and small on common lilac. 

In established plants, symptoms typically develop two or more years after infection.

EY can be mistaken for Dutch elm disease, especially by those not checking for 

EY. Symptoms of EY may include drooping and yellowing (fig. 37-3) usually in midto-late summer, leading to tree death within the next two years. The yellow leaves 

may drop or suddenly wilt, shrivel, turn brown, and remain attached to the twig. 

Roots are also affected, with fine roots dying first. The phloem and cambium change 

color from light cream to tan and then dark brown, typically as uniform discoloration, 

not streaking, although elongate dark brown flecks may be present. A faint wintergreen aroma emanates from freshly exposed, moist inner bark of infected branches, 

but this aroma is not always present and, when present, is difficult for many people 

to detect. In established trees, symptoms typically develop the season after infection. 
138 USDA Forest Service RMRS-GTR-335. 2016.

Figure 37-2—Lilac witches’-broom. (A) Late-season growth and deformed, scorched leaves of common lilac; (B) Multiple witches’-brooms at base of late lilac; (C) Late-season growth and late flowering 

lilac (James A. Walla, Northern Tree Specialties, used with permission).

A B C

Figure 37-1—Ash yellows witches’-brooms. (A) Several witches’-brooms on a single green ash trunk, 

with characteristic lime-green foliage; (B) Witches’ broom with dwarfed, deformed, and single leaflet 

leaves; (C) Witches’ broom in dormant season showing upright, narrow-angle branching and branching from leaf axils (James A. Walla, Northern Tree Specialties, used with permission).

A B C

Figure 37-3—Elm yellows symptoms. (A) Single American elm branch showing symptoms of wilted, 

yellowing leaves; (B) Witches’-broom on hybrid elm (James A. Walla, Northern Tree Specialties, used 

with permission).

A B
USDA Forest Service RMRS-GTR-335. 2016. 139

Symptoms commonly occur on the whole tree over a period of only a few weeks 

in mid- to late summer, although monitoring from early infection will often show 

symptoms spreading from one branch or portion of a tree. On slippery elm, similar 

symptoms develop, but small witches’-brooms also form just before a tree dies, the 

discolored bark is inconsistent, and the aroma when present is like maple syrup.

Disease Cycle

Vector insects typically transmit the pathogens while feeding on tree leaves in summer 

and early fall. The pathogens spread in the phloem tissues of infected plants where they 

overwinter primarily in roots. Pathogen survival may be reduced in aboveground parts in 

colder winters. In spring, phytoplasmas spread to greater portions of infected plants, and 

may be acquired by host insects feeding on the leaves. There is a latent period of three to 

four weeks before the phytoplasmas are transmitted to new hosts to complete the disease 

cycle. Phytoplasmas can also be transmitted by vegetative propagation. These pathogens 

are not known to be transmitted through seed.

Damage

AshY causes progressive growth suppression, reduced competitiveness, and decline 

in susceptible white and green ash, but there is wide variation in these effects. AshY and 

LWB reduce life expectancy and aesthetic value of susceptible hosts. Reduced pathogen 

aggressiveness, increased host tolerance, and good growing conditions typically result 

in reduced damage by AshY and LWB. A moderately susceptible tree may exhibit only a 

reduced growth rate, which may be desirable in some landscape settings. Decline resulting from these diseases may be triggered by occurrence of stress, whereby infected trees 

do not recover but noninfected trees do. Surveys of trees in six states and three provinces 

in the Great Plains did not find an association between tree condition and AshY phytoplasma presence in green ash, likely because there are multiple factors that damage ash. 

Individual ash and lilac have been severely affected by these diseases in North Dakota 

(fig. 37-4). Evaluation in North Dakota found very wide variation in both pathogen aggressiveness (fig. 37-5) and green ash tolerance.

In the primary range of EY, there are periodic disease outbreaks, and most North 

American elm species are killed in those areas. EY kills most North American elm species one or two years after infection. It has been observed that infected slippery elms may 

survive for several years. In the more tolerant Eurasian elm species, trees that have been 

inoculated grow slowly and may develop chlorosis and witches’-brooms. As EY affects 

Figure 37-4—Damage caused by ash yellows and lilac witches’-broom. (A) Reduced and sparse 

growth on green ash; the center tree was infected by an aggressive ash yellows phytoplasma isolate. 

(B) Reduced growth, sparse foliage, and deformed shape of late-blooming lilac (James A. Walla, 

Northern Tree Specialties, used with permission).

A B
140 USDA Forest Service RMRS-GTR-335. 2016.

elms, the trees attract and support large populations of the bark beetles that vector the 

Dutch elm disease fungus. It is not unusual for an elm to have both EY and Dutch elm 

disease and for Dutch elm disease to increase greatly in a location where EY is active. EY 

apparently occurs at a very low level of incidence in the Great Plains and is not known to 

have caused substantial damage there.

Management

The best disease management for AshY, LWB, and EY is use of tolerant species or 

selections. Green ash and velvet ash are more tolerant than is white ash to AshY. Three 

of six green ash cultivars (‘Bergeson,’ “Dakota Centennial®,” ‘Patmore’) and one of 

five white ash cultivars (‘Autumn Applause’) are tolerant of AshY. The late-blooming 

lilac species and hybrids are less tolerant to LWB damage, whereas common lilac is 

relatively tolerant. Eurasian elms are more tolerant than the native elm species, and they 

have not been damaged when exposed to natural infection in North America. ‘Frontier,’ 

‘Pathfinder,’ and ‘Patriot’ hybrid elms appear to be tolerant to EY, and “Homestead” 

hybrid elm appears to be resistant to EY. 

Calcium oxytetracycline is labeled as a systemic antibacterial for seasonal suppression of AshY, EY, and some other phytoplasma diseases. Injected trees remain infected, 

but temporary relief from damage may be provided for young infected trees. Disease 

prevention may be achieved if the antibacterial is used before or at the time of infection. 

Concern for development of microbial resistance to antibiotics should be considered 

before they are used for management of phytoplasma diseases. Damage to trees susceptible to some phytoplasmas is substantially reduced by grafting onto rootstocks tolerant 

to those phytoplasmas; such has not been reported for AshY, LWB, or EY, but would be 

expected to hold true. Management of AshY should be tempered by consideration of the 

long-term prospect of loss from emerald ash borer.

Selected References

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Walla, J.A.; Jacobi, W.R.; Tisserat, N.A.; Harrell, M.O.; Ball, J.J.; Neill, G.B.; Reynard, 

D.A.; Guo, Y.H.; Spiegel, L. 2000. Condition of green ash, incidence of ash yellows 

phytoplasmas, and their association in the Great Plains and Rocky Mountain regions 

of North America. Plant Disease. 84: 268–274.

Figure 37-5—Effects of ash yellows phytoplasma aggressiveness on growth of green ash seedlings. (A) Non-inoculated control; (B) Low aggressiveness; (C) Moderate aggressiveness; (D) High 

aggressiveness (James A. Walla, Northern Tree Specialties, used with permission).

A B C D
USDA Forest Service RMRS-GTR-335. 2016. 141

38. Armillaria Root Disease

James T. Blodgett; revised from Lloyd R. Fuller and Robert L. James 

(Riffle and Peterson 1986)

Armillaria root diseases are caused by several species of fungi in the genus Armillaria. 

These pathogens are common facultative parasites on more than 600 plant species 

throughout the world. Root diseases caused by these fungi are responsible for considerable economic damage in natural forests, plantations, orchards, and vineyards.

Hosts and Distribution

Root diseases caused by Armillaria species occur sporadically throughout the Great 

Plains, but their distribution in most of the Great Plains states is not well documented. 

Most trees, shrubs, vines, and some herbaceous plants are susceptible to these pathogens. 

Armillaria root diseases are common in windbreaks, forests, and landscapes. Some susceptible plants native to or planted in the Great Plains include: apples (Malus spp.), aspen 

(Populus spp.), boxelder (Acer negundo), cottonwoods (Populus spp.), oaks (Quercus

spp.), paper birch (Betula papyrifera), Peking cotoneaster (Cotoneaster acutifolius), 

ponderosa pine (Pinus ponderosa), poplars (Populus spp.), Scots pine (Pinus sylvestris), 

stone fruits (cherry, peach, and plum; Prunus spp.), Tatarian honeysuckle (Lonerica 

tatarica), willows (Salix spp.), and members of the rose family (Rosaceae).

Symptoms and Signs

Symptoms are not always diagnostic, and trees with Armillaria root disease do not 

always show well-defined symptoms. When present, crown symptoms resemble those of 

other root disorders: chlorotic foliage, reduced terminal growth, premature foliage drop, 

branch dieback, and stress crop of 

seeds or cones (fig. 38-1). One or 

more of these symptoms may be 

present on a single tree over many 

years or sudden crown mortality 

may occur. Sudden crown mortality 

can occur even if a tree was infected 

for several years. Hosts infected at 

the base of the trunk (root collar) 

may have external basal resinosis 

or gummosis (fig. 38-2), but these 

symptoms may not always be present. Armillaria species cause white 

rot, often with zone lines (fig. 38-3), 

in roots and lower stems. Decay can 

be difficult to detect unless trees are 

uprooted or break at the root collar.

Diagnostic signs include mycelial fans under or in the bark, 

rhizomorphs (a cordlike structure 

containing mycelia, which resembles roots or shoe strings), and 

mushrooms. Armillaria species 

produce characteristic mycelial fans 

(fig. 38-4) that appear as white to 

light cream-colored, fanlike mycelial mats beneath or inside the bark 

of roots and lower stems (butts). 

Figure 38-1—Chlorotic foliage with extensive dieback, 

a typical symptom of (A) lodgepole pine (Pinus contorta) 

and (B) aspen infected with Armillaria root disease 

(James T. Blodgett, U.S. Forest Service).

A

B
142 USDA Forest Service RMRS-GTR-335. 2016.

Figure 38-2—Lodgepole pine infected with Armillaria root disease 

showing resinosis at the root collar (James T. Blodgett, U.S. Forest 

Service).

Figure 38-3—Armillaria white rot with zone lines (Daniel H. Brown, U.S. 

Forest Service, Bugwood.org).

Figure 38-4—Mycelial fan, a diagnostic sign of Armillaria root disease, seen here 

on aspen (James T. Blodgett, U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 143

Mycelial fans develop in the 

cambial zone of roots and root 

collars, and can spread into stems. 

The rootlike rhizomorphs may be 

found on root surfaces, in adjacent 

soil (fig. 38-5), in decayed wood, 

or under the bark of living or dead 

trees. These structures are gray, 

reddish brown, or black on the 

exterior with a white inner core and 

tip. Rhizomorphs branch like roots, 

but typically have less variation in 

diameter (0.5 to 2 mm) compared 

to roots. Rhizomorphs that occur 

on root surfaces, in soil, and in decayed wood tend to be round; ones 

beneath bark tend to be flat.

Armillaria species might produce mushrooms in sporadic years 

(fig. 38-6). Mushrooms are shortlived, but may be present from late 

summer through fall if moisture is 

adequate. The mushrooms grow 

in small to large clusters on living 

and dead hosts, or on soil near 

underground wood. Armillaria

mushrooms can be identified by 

their honey-yellow caps 1 to 5 inches across with a persistent annulus 

(ringlike structure) on the upper 

stem. Caps have fine erect hairs or 

scales on the upper surface. The 

white gills on the underside of the 

cap are attached to the stem. Spores 

are white to light cream colored.

Differences among the Armillaria

species and host species result in 

variation among symptoms and 

signs, and Armillaria species are 

often somewhat host specialized. Armillaria can be identified to species by traditional 

morphology, mating types, cultural characteristics, and DNA sequences. Additional studies 

are needed to identify Armillaria species present in many areas of the Great Plains.

Disease Cycle

These pathogens spread primarily as vegetative mycelium by root contact, root grafts, 

or rhizomorphs. Rhizomorphs can grow considerable distances (up to 6.6 ft per year) 

through soil and initiate parasitic infections or saprotrophic colonization of roots or root 

collars. Armillaria species can occupy sites for many years. In one relatively dry region, a 

single vegetative clone of Armillaria was shown to occupy an area of approximately 

2,384 acres and was estimated to be up to 8,650 years old. During favorable late summer 

or fall weather, mushrooms of Armillaria species may produce prolific wind-dispersed 

spores. However, unlike many other decay pathogens, the spores rarely initiate disease.

Mycelial strands can colonize susceptible host cambium and initiate canker formation. Cankers can remain dormant (quiescent) or progress into lethal infections. Healthy, 

vigorous hosts or resistant host species may survive for many years. Stressed hosts, hosts 

with numerous or extensive infections, and susceptible host species can be killed quickly. 

Successfully infected hosts usually die when the fungus kills the cambium around root 

Figure 38-6—Fruiting bodies of an Armillaria species at 

the base of a paper birch (James T. Blodgett, U.S. Forest 

Service).

Figure 38-5—Rhizomorphs (gray to black, see arrows) 

in soil growing into a root and root collar, diagnostic signs 

of Armillaria root disease (James T. Blodgett, U.S. Forest 

Service).
144 USDA Forest Service RMRS-GTR-335. 2016.

collars. Live trees can be uprooted or break at the lower stem because of mechanical 

failure resulting from decay. Although such mechanical failures are more frequent during 

high-wind conditions, they can occur during conditions of low or no wind.

Saprotrophic colonization of wood tissues occurs after trees or tree tissues die. Armillaria

species can also colonize trees killed by other damage agents. The fungus can survive for 

decades in dead roots or stumps, and can infect new trees over the course of decades.

Damage

Armillaria root diseases affect trees used for many forest services and products including recreation, aesthetics, timber, windbreaks, watershed protection, soil stabilization, 

and carbon sequestration. Root diseases in developed sites can produce hazard trees that 

threaten life and property because of an increased probability of hosts falling. Armillaria 

root diseases cause direct tree mortality and indirect mortality through uprooting or 

breakage of lower stems, growth reduction, and wood loss from decay. They also predispose trees to other lethal agents including insects and other diseases. Armillaria root 

diseases are common contributing factors in many declines and can increase the likelihood of fire by increasing the accumulation of dead fuels.

In some situations, Armillaria species are not damaging unless hosts are under stress. 

Stresses can include extended droughts or dry sites, competition for light or nutrients, 

insect attacks, infections by other pathogens, or poor planting technique including offsite 

plantings. Young trees are often quickly killed by Armillaria species, whereas 15- to 

20-year-old trees tend to be more tolerant.

Management

These pathogens are difficult or impossible to eradicate. Management measures can 

include removing infected stumps and roots, limiting underground spread of the fungus, 

managing for resistant species, using proper planting techniques, and maintaining tree 

health. Management of root diseases over extensive acreage is seldom possible or practical. Valuable ornamental, shade, and orchard trees that are adjacent to infected trees 

may be protected somewhat by removal of infected stump and root material from the 

soil. Chemicals have been used to sanitize infested soil and wood, but only professional 

chemical applicators should apply these chemicals near healthy trees. Favoring healthy 

existing species and favoring species that are resistant to Armillaria root disease during 

thinning or planting offer the most promising and long-lasting approaches to disease 

control. Lists of resistant plants that are well adapted to a locale can be obtained from 

local extension educators or from the reference by Raabe and McCain (1967). Use proper 

planting methods, such as adequate site preparation, selection of species suitable for the 

site, use of foliage that is in balance with root systems, selection of disease-free planting 

stock, and prevention of J-roots (tap roots pointing up; usually the result of improper 

planting). Reduce host stress by thinning, weed control, watering during droughts, and 

managing other diseases and insect pests so that the impact of Armillaria root diseases 

can be minimized.

Selected References

Ferguson, B.A.; Dreisbach, T.A.; Parks, C.G.; Filip, G.M.; Schmit, C.L. 2003. Coarsescale population structure of pathogenic Armillaria species in a mixed-conifer forest 

in the Blue Mountains of northeast Oregon. Canadian Journal of Forest Research. 33: 

612–623.

Miller, O.K., Jr. 1972. Mushrooms of North America. New York: E.P. Dutton & Co. 360 p.

Raabe, R.D. 1962. Host lists of the root rot fungus Armillariella mellea. Hilgardia. 33: 25–88.

Raabe, R.D.; McCain, A.H. 1967. Plants resistant or susceptible to Armillaria root rot. 

Agricultural Extension Service Bulletin AXT-6. Davis, CA: University of California. 6 p.

Shaw, C.G., III; Kile, G.A. 1991. Armillaria root disease. Agric. Handb. 691. Washington, 

DC: U.S. Department of Agriculture, Forest Service. 233 p.

U.S. Department of Agriculture. 1960. Index of plant diseases in the United States. 

Agric. Handb. 165. Washington, DC: U.S. Department of Agriculture, Crops Research 

Division. 531 p.
USDA Forest Service RMRS-GTR-335. 2016. 145

39. Ganoderma Root Rot or White Mottled Rot

James T. Blodgett

Ganoderma root rot, also called white mottled rot, is caused by the fungus Ganoderma 

applanatum. This fungus is found in all 50 states and occurs throughout North America 

and Europe. It is a pathogen and a common wood-decaying fungus of roots and lower 

stems (butts) of many deciduous and some coniferous trees species. Ganoderma-caused 

root rot has been reported in live trees, such as apple (Malus spp.), aspen (Populus spp.), 

basswood (Tilia spp.), beech (Fagus spp.), birch (Betula spp.), cherry (Prunus spp.), 

citrus (Citrus spp.), cottonwood (Populus spp.), elm (Ulmus spp.), hemlock (Tsuga spp.), 

hornbeam (Carpinus caroliniana), horsechestnut (Aesculus hippocastanum), black 

locust (Robinia pseudoacacia) and honeylocust 

(Gleditsia triacanthos), maple (Acer spp.), mulberry (Morus spp.), oak (Quercus spp.), spruce 

(Picea spp.), sycamore (Platanus occidentalis), 

tulip tree (Liriodendron tulipifera), sweetgum 

(Liquidambar styraciflua), and willow (Salix

spp.).

G. applanatum is commonly known as the 

artist’s conk. The name comes from the use 

of its fruiting bodies as a drawing medium by 

artists (fig. 39-1). When the fresh lower surface 

is rubbed or scratched, it immediately changes 

from white to dark brown, producing shading or 

visible lines. When the conk is dried, drawings 

become permanent.

Hosts and Distribution

In the Great Plains, G. applanatum occurs predominantly in aspen, cottonwoods, and 

other Populus species. This fungal pathogen is irregularly distributed across the Great 

Plains, but its distribution is not well documented in many areas.

Symptoms and Signs

Crown symptoms in trees affected by G. applanatum are not always apparent. When 

present, symptoms resemble those of other root disorders, including reduced terminal 

growth, chlorotic foliage, premature foliage drop, and branch dieback. Although 

G. applanatum is often found in dead trees in an affected stand, it can decay roots and 

butts of otherwise healthy trees in the stand.

Extensive wood decay in roots and butts of infected trees is often overlooked until 

trees fall. Trees with this root disease often break at the soil line or slightly below with 

few or no attached roots (fig. 39-2). Often no other symptoms are apparent. Small trees 

or trees growing on poor sites with dry, shallow soils may be killed before decay leads to 

Figure 39-1—Drawing on the lower 

surface of Ganoderma applanatum conk 

(James J. Worrall, U.S. Forest Service).

 Figure 39-2—Roots infected with Ganoderma applanatum. Roots often break (A) near the stem or 

(B) at root collars. Conks might be present (A) above or (B) below the break (James T. Blodgett, U.S. 

Forest Service).

A B
146 USDA Forest Service RMRS-GTR-335. 2016.

windthrow. The fungus causes a white rot, and decayed wood is mottled white and light 

tan, eventually becomes spongy, and may develop black lines (fig. 39-3).

Fruiting bodies (conks) are the best indicator of host infection, but conks typically are 

not formed until several years after initial infection. Although conks are not produced on 

some trees with extensive decay, conks often develop before trees fall or die. Conks also 

can develop after trees fall or die.

G. applanatum typically produces firm, flat, perennial, shelflike conks at the base of 

trees, often near old wounds, and usually close to the soil (fig. 39-4). Conks can vary in 

appearance, starting as a small white projection from bark. The upper surface is initially 

white, changing to gray, beige, or brown with age. The lower surface and conk margin 

remain white when growing, and the lower surface is covered with fine pores. When the 

conk is cut, distinct mottled brown annual tube layers (hymenial tissues) can be observed 

(fig. 39-5). Context tissues (non-hymenial tissues that compose fungal fruiting bodies) are 

brown. When conks or parts of conks die, the smooth upper and lower surfaces become 

dark gray to black and develop cracks. Conks can grow to a diameter of more than 16 

inches, and both living and dead conks can persist for years.

Figure 39-4—Conks of Ganoderma applanatum. They can vary in appearance, but have a white 

margin and underside when growing (James T. Blodgett, U.S. Forest Service).

A B C

Figure 39-3—White mottled rot caused by Ganoderma applanatum (A-C), with (B, C) clear mottling, 

and (C) a zone line (right) (A: James T. Blodgett, U.S. Forest Service; B, C: James J. Worrall, U.S. 

Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 147

Disease Cycle

Infections occur from airborne spores or from contact 

with infected roots of neighboring trees. Conks can produce 

billions of airborne spores that 

are released from pores on the 

lower surface. Although not 

well understood, infections 

likely occur at wounds on roots 

or at the base of trees. Root-toroot spread can result in groups 

of infected trees called disease 

centers. In addition, the fungus 

can persist as a saprobe for several years after the host dies.

Damage

Ganoderma root rot affects trees used for many forest products and services including 

timber, recreation, and aesthetic value. It is more common in older trees and may be more 

prevalent on moist sites that are well suited for Populus species. Ganoderma root rot 

causes direct tree mortality, indirect tree mortality by breakage at the base or uprooting, 

growth reduction, wood loss from decay, and predisposition of trees to other disease and 

insect agents. Special consideration should be given when this disease occurs in developed sites because live, healthy-looking hosts with this disease frequently fall, which 

can endanger lives and property. Because the pathogen kills and decays roots, it likely 

reduces root suckering (vegetative formation of new stems from roots) and regeneration 

success.

Management

Once Ganoderma root rot is established in trees, practical methods for reducing this 

disease are unavailable. Because a relationship exists between wounds and decay, avoid 

wounds to reduce future infections. This disease is a major problem of aspen and other 

Populus species, and only a few other deciduous tree species in the Great Plains, so 

recognizing the problem and discriminating against susceptible host species is a practical 

management option. Species conversion could include complete removal of susceptible 

host tree species from an affected stand, or selective removal of aspen and other host species during thinning to favor nonhost species.

At developed sites with hosts, hazard tree inspectors should be trained to recognize 

the conks of G. applanatum. Conks are the only clear indicator of infection and extensive 

decay in live, standing trees. Fortunately, conks often develop before trees fall or die, providing an opportunity for removal of hazard trees. Species conversion away from hosts 

susceptible to Ganoderma root rot should be strongly encouraged on developed sites, 

especially if this disease is present.

Selected References

Hinds, T.E. 1985. Diseases. In: DeByle, N.V.; Winokur, R.P., eds. Aspen: ecology and 

management in the Western United States. Gen. Tech. Rep. RM-119. Fort Collins, CO: 

U.S. Department of Agriculture, Forest Service, Rocky Mountain Forest and Range 

Experiment Station: 87–106.

Ross, W.D. 1976. Fungi associated with root diseases of aspen in Wyoming. Canadian 

Journal of Botany. 54: 734–744.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Figure 39-5—Ganoderma applanatum produces perennial 

conks with mottled brown annual tube layers (hymenial tissues) and brown context tissues (James T. Blodgett, USDA 

Forest Service).
148 USDA Forest Service RMRS-GTR-335. 2016.

40. Nematodes of Broadleaf Trees

Timothy C. Todd and Jon A. Appel

Plant-parasitic nematodes have long been implicated as contributors to poor vigor and 

stunting of seedlings in forest-tree nurseries and decline of trees in windbreaks. These 

microscopic roundworms feed on root tissues by using a hollow needlelike mouthpart 

called a stylet (fig. 40-1). Although ubiquitous in agricultural and native soils, most plantparasitic nematodes become economically important only at large population densities 

or when the vulnerability of their host is amplified by developmental stage (especially 

seedling), environmental stress, or opportunistic pathogens. 

Hosts and Distribution

Information on nematode associations with specific tree species in the Great Plains 

is sparse. Windbreak surveys have been conducted in several states, but the presence of 

grasses and weed cover limit conclusions about host status in most cases. Nevertheless, 

surveys have shown that plant-parasitic nematodes are widespread in windbreaks in the 

Great Plains. Many of these species have broad host ranges, so it is likely that understory 

grasses and forbs contribute to conditions favorable for the buildup and maintenance of 

large population densities.

The dagger nematode Xiphinema americanum (fig. 40-1) is the most frequently 

encountered plant-parasitic nematode in Great Plains windbreak surveys (table 40-1). A 

survey of 76 South Dakota windbreaks and natural stands conducted during 1965–1967 

recovered the dagger nematode from nearly 90 percent of samples; large populations 

(>100 individuals/100 cm3 [6 cubic inches] soil) most often were associated with stands 

of eastern cottonwood (Populus deltoides), elm (Ulmus spp.), hackberry (Celtis occidentalis), and Russian-olive (Elaeagnus angustifolia). Dagger nematodes were less prevalent 

in a 2004-2006 survey1 of 81 Kansas windbreaks (43 percent of total survey samples; 

table 40-1), but relatively high frequencies of occurrence (60-64 percent) were observed 

for green ash (Fraxinus pennsylvanica) and hackberry stands (table 40-2). Dagger 

nematode population densities exceeding 100 individuals/100 cm3 soil most often were 

associated with these tree species.

1 Data on file with the Kansas Department of Agriculture, Topeka, KS.

Figure 40-1— Anterior of an adult female dagger nematode, Xiphinema 

americanum, showing the elongated odontostyle (Tim Todd, Kansas State 

University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 149

Table 40-1—Prevalence of nematode genera recovered from Great 

Plains windbreak surveys

Nematode common Survey (number of samples)

name (genus) South Dakota (113) Kansas (157)

- - - - - - - - - Percent positive- - - - - - - -

Dagger (Xiphinema) 88 43

Lance (Hoplolaimus) 33 <1

Pin (Paratylenchus) 75 54

Ring (Mesocriconema) 4 8

Root-lesion (Pratylenchus) 9 27

Spiral (Helicotylenchus) 45 58

Stunt (Tylenchorhynchus) 67 38

Stubby-root (Paratrichodorus) 7 4

Pin (Paratylenchus spp.), stunt (Tylenchorhynchus spp.), and spiral (Helicotylenchus spp.) 

nematodes also occur at high frequencies in Great Plains windbreaks (table 40-1). The 

largest population densities of spiral and pin nematodes in Kansas windbreaks are associated with hackberry and Russian-olive, respectively (table 40-2).

Plant-parasitic nematodes are widely distributed in forest nurseries as well and occasionally are associated with hardwood seedlings, particularly in the southern United 

States. Notable examples include the root-knot nematode Meloidogyne incognita on 

flowering dogwood (Cornus florida) and the needle nematode Longidorus americanus on 

white oak (Quercus alba). Although L. americanus damage has been observed only for 

pine (Pinus spp.) seedlings, several species of oak may serve as reservoir hosts, leading 

to severe stunting of loblolly pine (P. taeda) seedlings when grown in rotation.

Table 40-2—Prevalence and abundance of nematode genera recovered from four dominant tree species 

during a three-year survey of 81 Kansas windbreaks.

Prevalence Mean (maximum) number of

(percent) nematodes/100 cm3 soil

Nematode Russian Russian

common name Ash Elm Hackberry olive Ash Elm Hackberry olive

(genus) (n=15) (n=22) (n=11) (n=15) (n=15) (n=22) (n=11) (n=15)

Dagger 60 27 64 47 17 (160) 9 (110) 25 (160) 13 (70)

(Xiphinema)

Lance 0 5 0 0 0 (0) 1 (12) 0 (0) 0 (0)

(Hoplolaimus)

Pin 47 64 55 60 133 (900) 116 (930) 43 (320) 252 (2,210)

(Paratylenchus)

Ring 7 5 27 0 3 (40) 0 (10) 87 (590) 0 (0)

(Mesocriconema)

Root-lesion 20 14 18 47 17 (200) 3 (40) 4 (30) 18 (80)

(Pratylenchus) 

Spiral 87 68 91 20 129 (860) 122 (750) 272 (1,300) 8 (60)

(Helicotylenchus)

Stunt 40 50 55 33 7 (40) 14 (60) 38 (120) 45 (270)

(Tylenchorhynchus)

Stubby-root 7 14 0 0 0 (4) 2 (30) 0 (0) 0 (0)

(Paratrichodorus) 
150 USDA Forest Service RMRS-GTR-335. 2016.

Symptoms and Signs

Visual symptoms of nematode damage rarely are diagnostic. Aboveground symptoms 

of nematode damage to tree roots mimic drought stress and nutrient deficiencies, and 

include stunting, chlorosis, and a general lack of vigor. These symptoms can be extreme 

in seedlings. Damage to mature trees, in contrast, often is manifested as a slow decline. 

Belowground symptoms frequently are manifested as reductions in numbers of lateral 

and feeder roots, and may include damage that is unique to the nematode species present, 

such as galling, necrotic cortical lesions, and root tip injury (for example, swelling and 

necrosis).

A nematode assay of soil and roots collected from the drip line provides the best 

indication of a nematode problem; however, diagnosis can remain problematic, even with 

information on types and numbers of plant-parasitic nematodes present in the soil and 

roots. For perennial hosts, the expression of symptoms related to nematode feeding often 

is dependent on their interaction with other factors such as environmental stresses and 

other disease-causing organisms.

Disease Cycle

The life cycle of plant-parasitic nematodes consists of four juvenile stages and an adult 

stage, each separated by a molt of the cuticle. Species reproduce by laying eggs, and are 

either sexual, with males and females common, or parthenogenetic, with males rare or 

absent. The length of life cycle differs with species, but typically requires approximately 

1 month for completion.

Juveniles and adults generally are vermiform, but root-knot nematode adult females 

are swollen and pear shaped. Ectoparasitic species such as dagger, spiral, and stunt 

nematodes remain in the soil throughout their life cycle and feed by penetrating root cells 

with their stylet. In the case of the dagger nematode, an elongate stylet permits feeding on 

internal root tissues. Endoparasites such as root-knot and root-lesion nematodes burrow 

into roots and feed on cortical or vascular tissues. Root-lesion nematodes remain migratory, moving between roots and soil, whereas root-knot nematodes establish permanent 

feeding sites and become sedentary.

Damage

Nematode feeding damage varies according to feeding habit and the nature of the 

digestive enzymes produced but ultimately is associated with disruption of root function (water and nutrient absorption and translocation). Ectoparasitic species that feed on 

root surface tissues cause little visual damage except at very high population densities. 

Ectoparasites like dagger nematodes, which feed on internal root tissues, are capable of 

causing damage at lower population densities, particularly when feeding at or near root 

tips. Endoparasitic species cause extensive mechanical damage during their migration 

through root tissues, and often are implicated as partners in disease complexes. 

The potential for damage by X. americanum has been demonstrated for inoculated 

greenhouse-grown cottonwood and green ash seedlings, and for nursery-grown golden 

willow (Salix alba ‘Vitellina’) and green ash transplants in fumigated versus nonfumigated field soil. Growth rates were reduced in the presence of the nematode, and root 

systems were devoid of feeder roots. Additional evidence for pathogenicity comes from 

windbreak surveys. Dagger nematode populations exceeding 100 individuals/100 cm3

soil in South Dakota and Kansas windbreaks are frequently associated with poor tree 

health.

Other nematodes that may be associated with tree decline in Great Plains windbreaks 

include the root-lesion and spiral. Root-lesion nematode population densities were 

inversely related to tree health in the Kansas windbreak survey. Poor tree health was 

predicted as population densities approached 200 individuals/100 cm3 soil, a level of 

nematode pressure that was observed only for green ash. Little additional information 

is available about the impact of root-lesion nematodes on green ash, and an assessment 
USDA Forest Service RMRS-GTR-335. 2016. 151

of the relationship between tree health and nematode population densities within roots 

may be necessary for reliable estimates of potential damage. Evidence for a relationship 

between spiral nematodes and poor tree health in the Great Plains is likewise limited and 

restricted to hackberry. Spiral nematode population densities exceeding 1,000 individuals/100 cm3 soil were associated with declining stands of this tree species in Kansas.

Despite numerous reports of the occurrence of plant-parasitic nematodes in forest 

nurseries, few examples of damage to broadleaf species have been documented. The most 

serious risks for broadleaf seedling damage in Great Plains nurseries appear to be associated with dagger and root-knot nematodes. In addition to damage due directly to feeding, 

dagger nematodes are known to transmit several virus diseases of broadleaf trees (for 

example, tobacco and tomato ringspot viruses (family Secoviridae).

Management

Chemical control of plant-parasitic nematodes in windbreaks is economically unviable. Weed control in windbreaks may limit nematode population increases while 

reducing competition for soil moisture, resulting in healthier trees that can better tolerate 

the damage caused by nematodes. Nematode management in orchards, for instance, has 

shifted in recent years toward stress avoidance by using cultural practices such as irrigation and fertility maintenance that promote better root development.

Soil fumigation is a viable option for nematode management in nurseries, and most 

alternatives to methyl bromide provide adequate control. An integrated approach often 

is necessary, however, owing to the high cost of fumigation and the ability of nematode 

populations to rebound after treatment. Combining soil fumigation with practices such as 

crop rotation with nonhost cover crops and field fallowing can be an effective strategy for 

suppressing nematode populations in forest nurseries. The choice of cover crop depends 

on the target nematode species, and generalizations are difficult. Pearl millet (Pennisetum

glaucum), for which hybrids with resistance to several nematode species have been developed, is considered to be one of the best options for nematode control.

Selected References

Cram, M.M.; Fraedrich, S.W. 2005. Management options for control of a stunt and 

needle nematode in Southern forest nurseries. In: Dumroese, R.K.; Riley, L.E.; Landis, 

T.D., tech. coords. 2005. National proceedings: Forest and Conservation Nursery 

Associations—2004. Proc. RMRS-P-35. Fort Collins, CO: U.S. Department of 

Agriculture, Forest Service, Rocky Mountain Research Station: 46–50.

Cram, M.M.; Fraedrich, S.W. 2012. Nematode damage and management in North 

American forest nurseries. Tree Planters’ Notes. 55: 27–35.

Malek, R.B. 1968. The dagger nematode, Xiphinema americanum, associated with 

decline of shelterbelt trees in South Dakota. Plant Disease Reporter. 52: 795–798.

Malek, R.B.; Smolik, J.D. 1975. Effect of Xiphinema americanum on growth of 

shelterbelt trees. Plant Disease Reporter. 59: 144–148.

Ruehle, J.L. 1973. Nematodes and forest trees—types of damage to tree roots. Annual 

Review of Phytopathology. 11: 99–118.
152 USDA Forest Service RMRS-GTR-335. 2016.

41. Brown Spot of Pine

Judith O’Mara and Aaron A. Bergdahl

In the Great Plains, brown spot can be a serious problem in Scots pine (Pinus sylvestris) plantings. It is easily confused with another important pine disease, Dothistroma 

needle blight. Brown spot is also damaging on young longleaf pine (P. palustris) trees in 

the southern United States. The disease is also referred to as brown spot needle blight.

Hosts and Distribution

Brown spot is primarily an issue on Scots pines in windbreaks and Christmas tree 

plantations in the Great Plains. It is occasionally seen on ponderosa (P. ponderosa), mugo 

(P. mugo), and white pine (P. strobus) as well. Brown spot also occurs northward into 

Manitoba and Ontario. In the southeastern United States, brown spot can be a serious 

problem on longleaf pine. 

Symptoms and Signs

Initial symptoms for brown spot are yellow spots that turn reddish brown and eventually girdle the pine needle. These dark bands are surrounded by a yellow halo and 

frequently develop droplets of dried resin. 

Needle tissue beyond the band turns 

brown, producing a half-needle scorch 

(fig. 41-1). Infected needles eventually 

turn brown and drop off the tree. Heavy 

disease pressure can result in extensive 

needle scorch and branch dieback in the 

lower portion of the tree (fig. 41-2). 

Brown spot is caused by the fungus 

Mycosphaerella dearnessii (syn. Eruptio 

acicola and Scirrhia acicola). In the Great 

Plains, the conidial or asexual state of the 

fungus (Lecanosticta acicola) is found 

to be associated with the disease. In the 

southeastern United States, both stages of 

the fungus can be found on longleaf pine 

throughout the year. L. acicola spores (conidia) form in acervuli that erupt through 

dead needle tissue (fig. 41-3). The conidia 

are cylindrical (15 to 35 μm × 3 to 4 μm), 

multicelled, curved, and olive colored. 

In culture, the fungus grows slowly and 

produces a dark olive-green, gelatinous 

colony (fig. 41-4).

Brown spot symptoms can look similar 

to another pine disease, Dothistroma 

needle blight, caused by D. septosporum

(also referred to in earlier literature as 

Mycosphaerella pini and Schirria pini) 

and D. pini, for which no teleomorph has 

been found (see chapter 43).

 The conidia for both brown spot and 

Dothistroma needle blight are similar, the 

primary difference being that the conidia 

for Dothistroma are colorless. Scots pine 

is not susceptible to Dothistroma needle 

blight; ponderosa and mugo pines are 

susceptible to both diseases. 

Figure 41-1—Brown spot needle banding and 

half-needle scorch (Jill O’Donnell, Michigan State 

University, used with permission). 

Figure 41-2—Brown spot damage on Scots pine 

Christmas trees (Jill O’Donnell, Michigan State 

University, used with permission). 

Figure 41-3—Fruiting bodies (acervuli) on pine 

needles (Isabel Munck, U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 153

Figure 41-4—Left) Spores (conidia) of Lecanosticta acicola (Isabel Munck, 

U.S. Forest Service); (Right) Lecanosticta acicola in culture on malt agar (Kirk 

Broders, University of New Hampshire, used with permission).

Disease Cycle

In the Great Plains, brown spot overwinters as acervuli on dead needles attached to 

the tree or on the ground. Spore dispersal and infection takes place in late spring to early 

summer of the following year (fig. 41-5). Spread of the disease is through rainsplash, 

insects, and shearing tools. New needle growth is more susceptible to infection than 

older needles. In the central Great Plains symptoms start developing in late summer and 

continue through the fall. 

Brown spot is favored by warm, wet conditions. In southern states, longleaf pine 

infection occurs over a range of temperatures, but daytime temperatures of 86 °F and 

nighttime temperatures of 70 °F are optimal. Similar conditions for infection could be 

expected for pine species growing in the Great Plains. Infection is also favored by prolonged periods of needle wetness. Crowded plantings, tall grass, or heavy weed pressure 

can lead to poor air circulation, favoring conditions for disease development.

Figure 41-5—Disease cycle on Scots pine (David Haasser, North Dakota State University, 

used with permission, Adapted from Riffle and Peterson, 1986).
154 USDA Forest Service RMRS-GTR-335. 2016.

Damage

Occurrence of brown spot on Scots pine has been reduced in the southern Great Plains 

because of heavy disease pressure from pine wilt, which has destroyed many Scots pine 

plantings. Although it is less of a problem in landscape plantings, brown spot is still a 

serious problem in Christmas tree plantations where Scots pine is a staple species. The 

biggest challenge for Christmas tree growers is that the needle symptoms start to become 

apparent early in the fall before the harvest period for the Christmas season. In some 

cases, growers apply green paint to avoid a loss in tree sales during the holiday season. 

Management

Management for brown spot needle blight starts with selection. Pine plantings should 

not be established with only a single species or variety of pine. If planting Scots pine, 

keep in mind that short-needled Scots pines are more susceptible to brown spot than 

long-needled varieties. Prolonged leaf wetness can lead to greater disease severity, so 

improving airflow can help dry out the foliage and reduce disease pressure. For better 

airflow, use proper plant spacing, control weeds, and keep grass alleyways mowed. 

Removal of pine needles from around the tree may also help to reduce the amount of disease inoculum that overwinters. In Christmas tree plantations avoid shearing during wet 

conditions, as the disease can be spread on the shears. 

During wet years, crowded plantings may require the use of one to two fungicide applications. Make the first application when needles are one-half their full length. A second 

application may be necessary three to four weeks later depending on the residual period 

of the fungicide selected. 

Selected References

Kais, A.G. 1971. Dispersal of Scirrhia acicola spores in southern Mississippi. Plant 

Disease Reporter. 55: 309–311. 

McGrath, W.T.; Prey, A.J.; Morse, F.S. 1972. Brown spot needle blight of Scotch pine 

Christmas trees in Wisconsin: sporulation and control. Plant Disease Reporter. 56: 

99–102.

Nicholls, T.H.; Skilling, D.D.; Hudler, G.W. 1973. Scirrhia acicula in Scotch pine 

Christmas tree plantations. Plant Disease Reporter. 57: 55–59. 

Peterson, G.W. 1981. Pine and juniper diseases in the Great Plains. Gen. Tech. Rep. 

RM-86. Fort Collins, CO. U.S. Department of Agriculture, Forest Service, Rocky 

Mountain Forest and Range Experiment Station. 47 p.

Phelps, W.R.; Kais, A.G.; Nicholls, T.H. 1978. Brown-spot needle blight of pines. 

Forest Insect & Disease Leaflet 44. Formatted for the Internet September 1997. U.S. 

Department of Agriculture, Forest Service. http://www.na.fs.fed.us/spfo/pubs/fidls/

br_spot_pines/brown-spot.htm. (January 8, 2014).

Sinclair, W.A.; Lyon, H. 2005. Brown spot needle blight of pines. In: Diseases of trees 

and shrubs. 2nd ed. Ithaca, NY: Cornell University Press.
USDA Forest Service RMRS-GTR-335. 2016. 155

42. Cercospora Blight of Junipers

Glenn W. Peterson and David S. Wysong (Riffle and Peterson 1986) 

(updated by Ned Tisserat)

Junipers (Juniperus spp.) and other members of the Cupressaceae family are infected 

by two closely related needle-blighting fungi, Pseudocercospora juniperi and Passalora 

sequoiae.

Hosts and Distribution

Cercospora blight, caused by the fungus P. juniperi (formerly Cercospora sequoiae

var. juniperi), is a widespread needle disease of junipers in the eastern Great Plains. It is 

particularly common in the eastern third of Kansas and Nebraska, where relatively high 

summer humidity and frequent rains favor disease development. The disease is also common in the eastern and central United States. Rocky Mountain juniper (J. scopulorum) 

and its varieties are particularly susceptible to the disease. Eastern redcedar (J. virginiana) is less susceptible and Chinese juniper (J. chinensis) varieties are very resistant. 

A second fungus, P. sequoiae (formerly C. sequoiae), also causes a disease referred 

to as Cercospora blight in Arizona cypress (Cupressus arizonica), oriental arborvitae 

(Platycladus orientalis), and Chinese juniper and is found throughout the southeastern 

United States (including Texas and perhaps southern Oklahoma in the Great Plains) and 

from Pennsylvania to eastern Nebraska.

Symptoms and Signs

Symptoms of Cercospora needle 

blight first appear in late summer and 

fall on older, inner spur or scale needles 

located nearest the main tree trunk 

(fig. 42-1). Infected needles initially 

are bronze but later turn more dull 

red and drop from trees in October 

and November (earlier in the northern 

Great Plains). Commonly, all leaves of 

a branchlet are affected. Defoliation in 

succeeding years continues from the inner portion of the branch toward the tip, 

and from the bottom of the tree toward 

the top. Severely infected trees are open 

and spindly and may appear as if they 

had been scorched by a 

fire. Symptoms caused by 

P. sequoiae on Arizona and 

Leyland (x Hesperotropsis 

leylandii) cypress are 

similar.

P. juniperi forms small 

(140 μm diameter) fuzzy 

or hairy cushion-shaped 

fruiting bodies (sporodochia), easily visible with a 

hand lens, on dead needles 

(fig. 42-2). These sporodochia produce elongate, 

Figure 42-1—Damage to a Rocky Mountain 

juniper windbreak. Note thin foliage in the lower 

half of the tree crown (Mark O. Harrell, Nebraska 

Forest Service, used with permission).

Figure 42-2—Fruiting bodies of Pseudocercospora juniperi showing fuzzy gray appearance of spores (conidia and conidiophores) 

(Glenn Peterson, USDA Forest Service, retired).
156 USDA Forest Service RMRS-GTR-335. 2016.

multiseptate, light brown spores (conidia) measuring 42 to 57 μm × 2.5 to 3.0 μm (fig. 42-3). The 

sporodochia of P. sequoiae are 50 μm in diameter 

and also produce multiseptate, light brown conidia 

(30 to 60 μm × 5.0 to 5.5 μm) that are more tapered than P. juniperi.

Cercospora blight is readily distinguished from 

Phomopsis and Kabatina blights of junipers. The 

branches of Cercospora-infected trees usually will 

be devoid of foliage near their bases but will have 

healthy foliage on their tips, whereas branches of 

trees infected with Phomopsis and Kabatina blight 

will have dead needles at branch tips and healthy 

foliage near the branch base.

Disease Cycle

Spores of P. juniperi and P. sequoiae are dispersed by rainsplash from late April 

through October. There is little or no long-distance wind dispersal of spores; almost all 

spores are deposited within a few feet of the initial site of infection. Infection is more severe when precipitation during the growing season is above average. The period between 

initial infection and first appearance of symptoms is from two to three weeks.

Damage

Cercospora blight is more destructive than Kabatina or Phomopsis blights in established Rocky Mountain juniper plantings. Repeated defoliation weakens trees, reduces 

the functional and aesthetic value of windbreaks, and can result in tree mortality. 

Management

Rocky Mountain juniper should not be planted for windbreaks in much of the eastern 

Great Plains (east of longitude 98 degrees west) and where Cercospora blight is a problem because the disease is simply too difficult to manage in tightly spaced plantings. 

Instead, select eastern redcedar or Chinese junipers that have good resistance to this 

disease. Cercospora blight is less of a problem on Rocky Mountain juniper in landscape 

plantings. Nevertheless, the disease may occasionally develop. Proper tree spacing, which 

promotes good air movement and rapid drying of foliage, will inhibit blight development 

of both pathogens. 

Chemical control may be necessary during wet summers. Although symptoms appear 

in late summer to fall, fungal infection of needles actually occurs in summer. At least two 

fungicide applications (copper-based products and other fungicide types) in early June 

(earlier for Passalora-caused blight) and again in early July are needed. A third application in mid- to late July may be necessary during wet summers. Fungicide applications at 

other times are ineffective. The foliage must be thoroughly covered, especially the lower 

two-thirds of the tree crown. Yearly fungicide applications may not be needed once the 

disease is controlled. However, carefully inspect trees yearly to monitor for recurrence.

Selected References

Peterson, G.W. 1977. Control of juniper blight caused by Cercospora sequoiae juniperi. 

American Nurseryman. 145: 50–51.

Peterson, G.W. 1981. Pine and juniper diseases in the Great Plains. Gen. Tech. Rep. 

RM-86. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky 

Mountain Forest and Range Experiment Station. 47 p.

Peterson, G.W.; Wysong, D. 1968. Cercospora blight of junipers: damage and control. 

Plant Disease Reporter. 52: 361–362

Sutton, B.C.; Hodges, C.S. 1990. Revisions of Cercospora-like fungi on Juniperus and 

related conifers. Mycologia. 82: 313–325. 

Tisserat, N.A.; Pair, J.C. 1997. Susceptibility of selected juniper cultivars to cedar-apple 

rust, Kabatina tip blight, Cercospora needle blight and Botryosphaeria canker. Journal 

of Environmental Horticulture. 15: 160–163. 

Figure 42-3—Spores (conidia) of 

Pseudocercospora juniperi (James A. 

Walla, Northern Tree Specialties, used 

with permission).
USDA Forest Service RMRS-GTR-335. 2016. 157

43. Dothistroma Needle Blight of Pines

Aaron D. Bergdahl and Judith O’Mara; revised from Glenn W. 

Peterson and David S. Wysong (Riffle and Peterson 1986)

Dothistroma needle blight (DNB) is a common and serious fungal disease of native 

and exotic pines in windbreaks, Christmas tree plantations, and ornamental plantings 

throughout the Great Plains. This chapter uses fungal nomenclature for the pathogens 

causing DNB based on recent molecular work. Earlier work with these pathogens has not 

been based on these methods and should be interpreted with this in mind. There has been 

limited confirmation of DNB species by using molecular methods in North America. 

Hosts and Distribution

Of the commonly planted pine species in the Great Plains, primary hosts of DNB 

are Austrian (Pinus nigra), ponderosa (P. ponderosa), and mugo (P. mugo) pines. Scots 

pine (P. sylvestris) is highly resistant to DNB. Dothistroma species have been recorded 

on more than 35 pine species and hybrids in the United States. D. pini, which is thought 

to be the most prevalent species causing DNB in the Great Plains, has been confirmed 

on whitebark (P. albicaulis), Swiss stone (P. cembra), limber (P. flexilis), Austrian, and

ponderosa pines. D. septosporum (also referred to in earlier literature as Mycosphaerella

pini and Schirria pini) is found in states neighboring the Great Plains, and thus it is not 

unreasonable to suspect that it also occurs in the Great Plains.

Symptoms and Signs 

Early symptoms of infection, seen in early fall, are resin-soaked, dark green bands and 

dark green, yellow, and tan spots on needles (fig. 43-1, top). Within one to two weeks, 

the bands and spots turn brown to reddish brown (fig. 43-1, middle). In the Great Plains, 

D. pini does not develop bands that are as numerous and as red as those on pines west of 

the Rocky Mountains, where the disease is referred to as red band disease and is likely 

caused by D. septosporum. On the most susceptible trees, the ends of the needles distal 

to the bands may die (fig. 43-2) during the fall after infection. On others, the needle ends 

Figure 43-1—Symptoms of infection by D. pini: (Top) Dark green banding, an 

early symptom of infection; (Middle) Reddish spotting at the point of infection as 

symptoms develop in the late season; (Bottom) A fruiting structure (acervulus) 

erupting through the epidermis (Glenn Peterson, USDA Forest Service, retired).
158 USDA Forest Service RMRS-GTR-335. 2016.

may die the next season or in some 

cases may not die. Within one year after showing dieback, the whole needle 

typically dies and is shed. Symptoms 

on most trees occur in the lower portion of the tree on the interior needles 

(fig. 43-3). Heavily infected trees are 

often highly susceptible genotypes or 

trees grown in tight spacing and with 

poor air circulation. 

Disease Cycle

Stromatic acervuli, in which 

asexual spores (conidia) are formed, 

develop first near the original infection site, but may later form in any 

portion of discolored needles. In 

Nebraska, some acervuli develop 

and mature sufficiently in the fall 

after infection to raise and split the 

needle epidermis (fig. 43-1, bottom), 

and more become visible the next 

spring. Acervuli develop and mature 

earlier in the southern Great Plains, 

allowing for same-season infection 

if adequate moisture occurs in the 

fall. Acervuli develop more slowly 

and are not visible in the fall of the 

infection year in the northern Great 

Plains. A flap of tissue usually covers 

acervuli as they rupture the epidermis 

of the needle. Conidia of D. pini are 

three- to five-septate and on average 

30.0 μm × 3.5 μm in vivo. Conidia 

of D. septosporum are three- to fiveseptate and on average 28 μm × 2 μm 

in vivo. Owing to a large overlap in 

spore sizes, this characteristic is not 

sufficient for differentiation between 

D. pini and D. septosporum.

The conidia (fig. 43-4), which are 

exposed as the epidermis is raised, 

are dispersed by rainsplash impacting 

acervuli any time during the growing 

season. New infections can occur on 

any age-class of susceptible needles 

during periods of wet weather or 

high relative humidity from May to 

October. 

Two growing seasons are required 

for completion of the life cycle during most years in most areas of the United States (in 

California and Oregon, what is likely D. septosporum, may complete a cycle in one year). 

In North America, the sexual stage of what is likely D. septosporum, has been found only 

in Alaska, British Columbia, California, and Oregon. The stromata of the sexual stage 

produce ascospores, whose role in the development of epidemics is not known. In contrast, much is known about the role of conidia in disease development.

Figure 43-2—Second-year ponderosa pine needles 

with Dothistroma needle blight symptoms (banding, 

partial scorch) on a tree from a site where only D. pini

was confirmed by molecular methods (James A. Walla, 

Northern Tree Specialties, used with permission).

Figure 43-3—Ponderosa pine with severe Dothistroma 

needle blight symptoms starting in lower and interior 

portions of the crown leading to tip dieback, needle 

loss, and a sparse lower crown. Molecular analysis of 

isolates from multiple trees in the planting found only 

D. pini (James A. Walla, Northern Tree Specialties, 

used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 159

Damage

DNB is a serious foliar disease 

on a wide range of pine species in 

the United States and is a pest of 

economic significance in plantation 

forestry in other parts of the world. 

The potential for damage by DNB 

is reduced in the Great Plains by 

relatively dry conditions, especially 

to the south, and short, cool growing 

seasons to the north. Still, successive 

years of severe infection in the Great 

Plains result in decreased growth, 

poor vigor and, sometimes, death. The 

disease makes pines in the landscape 

unsightly and Christmas tree and 

nursery landscape trees unmarketable, 

and pine trees planted for windbreak 

protection or privacy screens lose 

their effectiveness (fig. 43-3). 

Premature defoliation caused by DNB has been documented to halt planting of Austrian 

pine for Christmas trees and has resulted in complete failure of ponderosa pine plantings 

in states east of the Great Plains. DNB sporadically occurs and may damage natural 

stands of limber, lodgepole (P. contorta), and ponderosa pines in the western United 

States, including Montana and South Dakota.

Management

Copper-based and some other fungicides effectively prevent infection by Dothistroma

species. Research in Nebraska showed that fungicides applied twice in the growing 

season (fig. 43-5) have provided dependable control of DNB on Austrian and ponderosa 

pines: (1) An application of copper-based fungicide around the time of budbreak 

(mid-April in Kansas to mid-May in more northern parts of the Great Plains) protects 

second-year and older needles. Sporulation has been observed to occur considerably later 

in northern North Dakota (as late as mid-June to early July), highlighting the importance 

of local conditions when determining treatment time. (2) A second application about 

one month later protects current-year needles around the time needle susceptibility develops (current-year needles initially resist infection and do not become susceptible until 

several weeks after complete needle emergence, around July in Nebraska).

In other species of pines where current-year needles are not susceptible to DNB, a 

single application as needles complete emergence (for example, mid-May in Nebraska) 

will control the disease. A single fungicide application made after considerable needle 

Figure 43-4—D. pini (A) spores (James A. Walla, 

Northern Tree Specialties, used with permission), and 

(B) axenic culture. Culture was grown on Dothistroma 

sporulating media (20 g malt extract, 5 g yeast extract, 

15 g agar technical no. 3, 1000 ml distilled water; autoclave for 15 min at 121 °C/15 lb psi) (Irene Barnes, 

University of Pretoria, Pretoria, South Africa, used with 

permission). 

Figure 43-5—Dothistroma pini disease development schedule for the Great Plains. Note that phenology may be altered due to the geographic location of the host. Timing may vary due to weather and 

latitude (Altered from Peterson 1981). 
160 USDA Forest Service RMRS-GTR-335. 2016.

growth in late June or July has been used effectively by Christmas tree growers in the 

central United States. There is some risk in the single fungicide application procedure in 

southerly regions because infection could occur in previous years’ needles prior to the 

single early-season fungicide application.

If substantial infection occurs during a year in which fungicide has not been applied, a 

well-timed fungicide application the next year will provide good control. If little or no infection occurs in the year fungicide was not applied, spraying can be skipped for another 

year. Those managing high-value trees are advised to spray annually if DNB is present in 

the area. However, annual spraying may not be necessary in park, residential, and similar 

types of plantings where disease pressure may be lower and exist at a tolerable level.

Removing and disposing of needles from underneath trees will have little effect on 

DNB because infected needles persist on the tree, many shed needles get caught in the 

tree, and most infections occur in spring before needles dispersing inoculum are shed in 

the Great Plains. 

Work by Peterson and others in Nebraska in the 1970s and 1980s identified foreign 

and domestic resistant seed sources of Austrian and ponderosa pines. Needles of all ages 

were highly resistant in some of these seed sources, as compared to standard seed sources 

in which current-year needles become susceptible in July and older needles are susceptible. These tree improvement efforts were completed, although the extent to which trees 

from these sources exist in the Great Plains is unknown. Nurseries in the Great Plains that 

previously propagated the resistant seed sources were not doing so in 2010.

Selected References

Barnes, I.; Crouse, P.W.; Wingfield, B.D.; Wingfield, M.J. 2004. Multigene phylogenies 

reveal that red band needle blight of Pinus is caused by two distinct species of 

Dothistroma, D. septosporum and D. pini. Studies in Mycology. 50(2): 551–565.

Barnes, I.; Walla, J.A.; Bergdahl, A.; Wingfield, M.J. 2014. Four new host and three new 

state records of Dothistroma needle blight caused by Dothistroma pini in the U.S.A. 

Plant Disease. 98(10): 1443.1.

Peterson, G.W. 1967. Dothistroma needle blight of Austrian and ponderosa pines: 

epidemiology and control. Phytopathology. 57: 437–441.

Peterson, G.W. 1981. Pine and juniper diseases in the Great Plains. Gen. Tech. Rep. 

RM-86. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky 

Mountain Forest and Range Experimental Station. 47 p.

Peterson, G.W. 1984. Resistance to Dothistroma pini within geographic seed sources 

of Pinus ponderosa. Phytopathology. 74: 956–960.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.
USDA Forest Service RMRS-GTR-335. 2016. 161

44. Kabatina Blight of Juniper 

Andrea Ostrofsky and Glenn W. Peterson (Riffle and Peterson 1986) 

(updated by Ned Tisserat)

Eastern redcedar (Juniperus virginiana) and Rocky Mountain juniper (J. scopulorum), 

native to the Great Plains, are important trees in windbreak, wildlife, and landscape plantings. Branch tips of both species have been damaged by the fungus Kabatina juniperi.

Hosts and Distribution

Blight caused by the fungus K. juniperi is widespread in North America and especially 

in windbreak and ornamental plantings of eastern redcedar and Rocky Mountain juniper 

in the Great Plains. The fungus may also be found in ornamental plantings of Chinese 

(J. chinensis), savin (J. sabina), and creeping (J. horizontalis) junipers. 

Symptoms and Signs

Kabatina blight symptoms typically appear in late winter or early spring a few weeks 

before new growth resumes and when foliage changes from its dull green winter coloration to a darker green color of summer. The terminal two to six inches of diseased 

branches scattered throughout the tree or shrub first turn dull green, then red or yellow 

(fig. 44-1). Small ash-gray to silver lesions dotted with small black fruiting bodies (acervuli) of the fungus are visible at 

the base of the discolored tissue 

(fig. 44-2). The brown, desiccated twigs eventually break 

off at the lesion, usually shortly 

after new growth resumes but 

sometimes up to a year after discoloration. Blighting is restricted 

to the branch tips and usually 

does not cause extensive branch 

dieback or tree death except 

in highly susceptible plants in 

settings that favor severe disease 

development. Nevertheless, 

yearly tip dieback may result in 

abnormal bunching of the foliage 

(witches’-broom) and stunting of 

young trees or shrubs. 

The black, rounded to oval 

fungal acervuli, visible with a 

hand lens, develop in the epidermis beginning a few weeks 

after symptoms appear in spring and become more 

numerous the next month. Their numbers then decrease throughout the summer partly because some 

of the dead foliage is shed. The colorless (hyaline), 

unicellular, ellipsoid spores (conidia) measuring 

4.5 to 8.0 μm × 2.3 to 3.0 μm are produced successively at the tips of tapered spore-bearing cells 

that cover the surface of the acervulus, giving it a 

granular appearance (figs. 44-3 and 44-4). 

A second fungus, Sydowia polyspora (anamorph 

Sclerophoma pithyophila), causes similar symptoms at about the same time of year on junipers 

in other regions of North America and also 

occurs on several other conifers. A third fungus, 

Figure 44-1—Branches 

of Rocky Mountain juniper 

infected with Kabatina juniperi. Note the discoloration 

and dieback of the branch 

tips (Ned Tisserat, Colorado 

State University, used with 

permission).

Figure 44-2—Lesion 

with black fruiting bodies 

(acervuli) on branchlet of 

Rocky Mountain juniper 

(Ned Tisserat, Colorado 

State University, used with 

permission).

Figure 44-3—Fruiting bodies (acervuli) 

of Kabatina juniperi that have broken 

through the host epidermis. Note the 

granular appearance of the surface 

(James A. Walla, Northern Tree 

Specialties, used with permission).
162 USDA Forest Service RMRS-GTR-335. 2016.

Phomopsis juniperovora, also causes twig blighting 

although symptoms of this disease are most apparent in summer and may continue through the fall. 

Unlike Phomopsis blight, foliar blighting caused 

by K. juniperi occurs only before new growth appears and does not continue to progress through the 

summer. Microscopic identification is necessary to 

differentiate K. juniperi, S. pithyophila, and P. juniperovora. Conidia of K. juniperi and S. pithyophila

are similar to alpha conidia of P. juniperovora, but 

they do not contain refractive oil droplets. Conidia 

of S. pithyophila and K. juniperi are similar in 

size; however, those of S. pithyophila develop in 

pycnidia, whereas those of K. juniperi develop in 

acervuli. The prevalence of S. pithyophila in the Great Plains has not been determined, 

although if present it likely is not as widespread or common as K. juniperi.

Disease Cycle

The disease cycle of Kabatina blight is not completely understood. The fungus apparently enters plants through wounds. Many insects, including larvae of species of 

juniper webworm (Dichomeris sp.) and juniper midge (Contarinia sp.), are known to 

feed on juniper foliage and twiglets. Lesions of Kabatina blight frequently develop at 

these feeding sites. Because disease symptoms develop in late winter or spring as the 

foliage color changes, it is likely infection of wounded tissue occurs sometime relatively 

late in the previous growing season or perhaps after freeze damage in late fall. However, 

the precise period of infection is not known. The fungus, which can survive cold temperatures, probably overwinters within the branchlets.

Damage and Management

Kabatina blight rarely causes serious damage to mature trees in windbreak or farmstead plantings and management is not required. Control in landscape plantings may 

be desirable for aesthetic reasons, but few management practices have been developed. 

Control of wounding agents such as insects may reduce infection levels. Pruning and destruction of infected branchlets in late summer may reduce levels of inoculum, but could 

result in more wounded tissue that is susceptible to infection. The unsightly appearance 

of infected trees is often improved when lower branches elongate and the dead branch 

tips fall from the tree. Rocky Mountain juniper varieties are more susceptible to Kabatina 

blight damage than eastern redcedar. Chinese juniper is very resistant to the disease. 

Many juniper cultivars are reported to have good resistance; plant them in high-risk 

landscape areas.

Selected References

Hartman, J.R.; Doney, J.; Johnson, M.; Fountain, W.; McNiel, R. 1995. Evaluation of 

juniper cultivars for susceptibility to Kabatina tip blight, 1994. Biological and Cultural 

Tests for Control of Plant Disease. 44: 109–112.

Hsiang, T.; Huang, J.; Yang, L.; Cook, S.; Shen, Y.; Richter, K.J. 2000. Occurrence of 

Kabatina juniperi in Ontario and genetic analysis using RAPD markers. Canadian 

Journal of Plant Pathology. 22: 79–88. 

Ostrofsky, A.; Peterson, G.W. 1977. Occurrence of Kabatina juniperi on Juniperus 

virginiana in eastern Nebraska. Plant Disease Reporter. 61: 512–513.

Ostrofsky, A.; Peterson, G.W. 1981. Etiologic and cultural studies of Kabatina juniperi. 

Plant Disease. 65: 908-910.

Perry, R.G.; Peterson, J. L. 1982. Susceptibility and response of juniper species to 

Kabatina juniperi infection in New Jersey. Plant Disease. 66: 1189–1191.

Tisserat, N.A.; Pair, J.C. 1997. Susceptibility of selected juniper cultivars to cedar-apple 

rust, Kabatina tip blight, Cercospora needle blight and Botryosphaeria canker. Journal 

of Environmental Horticulture. 15: 160–163. 

Figure 44-4—One-celled spores of 

Kabatina juniperi, which form on the 

surface of the fruiting body (acervulus) (James A. Walla, Northern Tree 

Specialties, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 163

45. Lirula Needle Blight of Spruce

James A. Walla

The fungus Lirula macrospora causes needle blight of spruce (Picea spp.) across the 

Northern Hemisphere. As currently described, it likely consists of a complex of several 

species, with significant variation in morphology, regional occurrence, hosts, and life 

cycles. Damage caused by and management of the individual components of the L. macrospora species complex are similar if differences in length of the life cycle are taken into 

account.

Hosts and Distribution

In North America, L. macrospora has been reported on the native black (P. mariana), 

blue (P. pungens), Engelmann (P. engelmannii), red (P. rubens), Sitka (P. sitchensis), and 

white (P. glauca) spruce and the exotic Norway (P. abies) spruce. These represent the 

majority of diversity among spruce taxa. In the Great Plains, it has been found on white 

spruce (primarily Black Hills seed sources) and blue spruce.

L. macrospora has been found in all Canadian Provinces and Territories and at least 14 

States, including the Great Plains states of North Dakota, South Dakota, Wyoming, and 

Colorado. It occurs sporadically, with incidence typically at only a small portion of sites, 

but with wide variation in incidence and severity within sites and over time. The disease 

is most common in areas with relatively cool temperatures and greater precipitation. 

Symptoms and Signs

Only one component of the L. macrospora complex (Taxon A in Walla [1998]) has been 

found in the Great Plains. The remainder of this chapter relates specifically to that component.

The first symptom (yellow bands) appears on needles about 14 months after infection 

(August to October in North Dakota). The bands gradually become light purplish brown 

and expand over the entire needle by about 22 months after infection (April to May 

the next spring in North Dakota). The needles become reddish brown the same spring 

(fig. 45-1), and, by then, a characteristic black basal band has developed in the abscission layer at the base of the needle (fig. 45-2). Inconspicuous asexual spore-producing 

structures (pycnidia) develop on third-year needles about 22 months after infection (April 

in North Dakota). Large, black, smooth sexual spore-producing structures (hysterothecia) 

develop mostly on the lower surfaces (fig. 45-3) along the length of the needles about 

23 months after infection, within one month after pycnidia develop. Hysterothecia are 

easily visible except when they are just appearing. They are nearly as wide as the needle 

surface and can be various lengths, ranging up to the entire length of the needle (figs. 45-2 

and 45-3). Additional black bands develop around some needles (figs. 45-2 and 45-3) 

when hysterothecia appear. Infected needles slowly fade to tan (fig. 45-1) by about 

34 months after infection. When mature, hysterothecia have a lengthwise slit, which 

Figure 45-1—Discolored needles caused by Lirula macrospora. (A) Symptoms in spring: light infection of second-year needles, mixed infection of third-year needles, gray fourth-year needles with some 

broken; (B) Symptoms in fall: new symptoms on second-year needles, light brown discoloration of 

third-year needles (James A. Walla, Northern Tree Specialties, used with permission).
164 USDA Forest Service RMRS-GTR-335. 2016.

Figure 45-2—Mature Lirula macrospora fruiting bodies (hysterothecia), bands, asci, and spores: (A) 

Closed hysterothecia along length of needles, bands around needles, and basal bands; (B) Open 

hysterothecia, exposing ascus-bearing surface; (C) Ascus and ascospores stained in aniline-blue 

(James A. Walla, Northern Tree Specialties, used with permission).

A B C

Figure 45-3—Third-year symptoms and signs 

of Lirula macrospora on (A) upper, and (B) 

lower surfaces of the same needles. Black 

bands are visible on both surfaces. On these 

needles, fruiting bodies (hysterothecia) are 

visible only on the lower surfaces (James A. 

Walla, Northern Tree Specialties, used with 

permission).

B

A

opens (fig. 45-2B) to release ascospores (115 

to 160 μm × 2.5 to 3.5 μm) when the needles 

are wet. About 40 months after infection 

(September to October in North Dakota), 

the needles with mature hysterothecia 

shrivel and turn gray, but remain attached to 

branches until they are weathered off, typically another one or more years (fig. 45-1). 

The same three-year pattern of symptom and 

sign development occurs if older needles are 

infected, potentially resulting in a mixture of 

symptoms and signs on a single age-class of 

needles (fig. 45-1).

Symptoms, signs, and damage caused by 

Lirula brevispora and Isthmiella crepidiformis are similar to those caused by L. 

macrospora types that have a two-year life 

cycle. L. macrospora has often been misdiagnosed as Lophodermium piceae despite the 

distinctive symptoms, signs, and damage.

Disease Cycle

Lirula needle blight in North Dakota 

consistently has a three-year life cycle, as did 

samples observed from Manitoba, Minnesota, 

and South Dakota. Most infection occurs 

in the two months after budbreak. First 

symptoms develop about 14 months after 

infection. Pycnidia develop about 22 months 

after infection, and likely have an outcrossing role. Hysterothecia develop about 23 months after infection. Hysterothecia mature 

about one year after they first appear (three years after infection). Ascospore release from 

mature hysterothecia usually occurs during weeks with rain from just before bud-break 

through summer (late May through early September in North Dakota), with the greatest 

releases from June into August. The ascospores cause new infections to complete the 

disease cycle.
USDA Forest Service RMRS-GTR-335. 2016. 165

Damage

Damage by Lirula needle blight results 

from needle discoloration (aesthetic damage) 

and the loss of functioning needles (growth 

loss, death of branches). Trees with severe 

infection for a few years will exhibit a gradual 

reduction in length of new shoots throughout 

the crown, but especially on the most seriously affected branches, and if severe enough, 

those branches will stop growing and die. 

Because infected needles are not cast, severely 

infected trees appear to have a dirty green 

appearance from a distance (fig. 45-4), rather 

than the sparse appearance caused by needle 

casts. Long-term damage occurs only if lower 

branches were killed or if reduced growth 

allows competing trees to gain an advantage. 

Even when left unmanaged, Lirula needle 

blight is often not severe at sites in the Great 

Plains for more than a few years; reasons 

remain unclear.

Management

Lirula needle blight is poorly dispersed 

among sites, so great emphasis should be 

placed on avoiding introduction on transplants. 

Spruce should not be transplanted to a site 

where Lirula needle blight already occurs. 

There are varying levels of disease resistance 

among trees within species. Where some trees 

are severely infected, consider removing those 

trees to reduce disease pressure on neighboring spruce trees. In mixed plantings, favor 

retention of blue spruce over white spruce. 

Cultural actions that reduce the length of time 

that moisture remains on needles and that 

increase the distance between spruce trees will reduce potential disease severity. Until 

severe infection has occurred at least two years in a row, wait to see if it will subside 

on its own. If the disease has been severe (greater than 50 percent of two needle ageclasses infected) for more than two years, fungicide applications would be appropriate. 

Protection of needles for two months after budbreak in each of three consecutive years 

readily controls damage from Lirula. No fungicides are labeled specifically for Lirula 

needle blight. Some products containing azoxystrobin, chlorothalonil, copper hydroxide, 

copper salts of fatty and resin acids, copper sulfate, mancozeb, mono- and di-potassium 

salts of phosphoric acid, pyraclostrobin, sulfur, and thiophanate-methyl are labeled for 

use on spruce and have been found to control some foliar diseases. For unknown reasons, 

application of chlorothalonil in late July, well after initial spore release, provided the 

same excellent protection as multiple applications per season.

Selected References

Walla, J.A. 1995. Studies of Lirula macrospora (Hartig) Darker on spruce. Fargo, ND: 

North Dakota State University. 248 p. Ph.D. dissertation.

Walla, J.A. 1998. Variation in morphology of Lirula on Picea in North America. In: 

Laflamme, G.; Berube, J.A.; Hamelin, R.C., eds. Foliage, shoot and stem diseases of 

trees. Proc. IUFRO WP7.02.02 meeting. Information Report LAU-X-122. Sainte-Foy, 

QC: Canadian Forest Service: 6–12.

Figure 45-4—Heavy Lirula macrospora infection causing dirty-green appearance in (A) 

center tree, and (B) tree on right (James A. 

Walla, Northern Tree Specialties, used with 

permission).

A

B
166 USDA Forest Service RMRS-GTR-335. 2016.

46. Cyclaneusma Needle Cast of Pines

James A. Walla; revised from Glenn W. Peterson and James A. Walla 

(Riffle and Peterson 1986)

Needle cast caused by Cyclaneusma minus (syn. Naemacyclus minor) is commonly 

present on pines (Pinus spp.) and occasionally damages Scots pines (P. sylvestris) in 

young plantings in the Great Plains. A similar species, C. niveum, is present in North 

America, but has not caused damage.

Hosts and Distribution

Cyclaneusma minus infects many pine species, including Scots, Austrian (P. nigra), 

mugo (P. mugo), and ponderosa (P. ponderosa) pines. It occurs throughout the Northern 

Hemisphere. Most Scots pine plantings in North Dakota are infected, as well as young 

plantings in South Dakota, Nebraska, Kansas, and likely Colorado. It also occurs on 

mugo and ponderosa pine in the Great Plains.

Symptoms and Signs

In the eastern and central United States, where much research on C. minus has been 

done, infected needles typically become necrotic from September to November of their 

second growing season, with some infected needles becoming necrotic the next spring, 

summer, or fall. In the northern Great Plains, symptoms typically develop in September 

to October on third-year or older needles (fig. 46-1). The first symptoms are small 

light-green spots which gradually lighten and coalesce, turning the entire needle a characteristic dusty yellow or tan (fig. 46-2). Distinct transverse brown bars often occur on 

necrotic Scots pine needles. Needles are often cast two to three months after becoming 

fully discolored, but may be retained for several more months. Fruiting bodies (apothecia) 0.2 to 0.6 mm in length develop on necrotic needles (fig. 46-3A). Portions of needle 

tissue with apothecia will become lighter than surrounding tissue with age. Apothecia 

have slightly raised needle tissue over them, and are initially 

inconspicuous when closed, but 

the covers become lighter tan 

than the rest of the needle with 

age and are often conspicuous 

on recently cast needles and 

on damp needles. Apothecia 

are distinctive when they swell 

during wet weather and the 

covers swing open to reveal a 

dull cream to yellow hymenium 

(fig. 46-3B). Ascospores are 

clear (hyaline), two-septate, 

filiform, 65 to 100 μm long, 

and usually slightly bent like a 

boomerang.

Symptoms of this disease may 

be confused with natural and 

stress-related needle senescence 

or damage caused by aphids. 

The timing of C. minus symptom 

development and natural needle 

senescence overlaps, so needle 

symptoms must be used to 

Figure 46-1—Scots pine with third-year needles discolored 

by Cyclaneusma needle cast (James A. Walla, Northern Tree 

Specialties, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 167

differentiate the cause of necrosis. Naturally senescing Scots pine needles are bright yellow, and aphid damage may cause brown bars to occur on these needles. The presence 

of brown bars on dusty yellow or tan needles and presence of apothecia are indicative of 

Cyclaneusma needle cast. 

Disease Cycle

Spores are released and infection can occur throughout the year when temperatures 

are above freezing. Heavy infection can occur whenever spores are released, but tends 

to be greatest in spring before budbreak. All ages of needles are susceptible, beginning 

as soon as needles emerge from the fascicle sheaths. Older needles on susceptible trees 

develop symptoms as soon as two months after infection, and symptoms develop as soon 

as 15 months after infection of current-year needles. Infected needles of less susceptible 

trees remain symptomless for 12 to 24 months longer. Apothecia develop on attached or 

cast needles one to five months after first symptoms appear. They open when the needles 

are wet, and spores are dispersed from them by wind. The disease should be managed as 

if it has a two-year life cycle, although time from infection to spore production on older 

needles can be less than one year. 

Damage

This fungus has caused extensive early yellowing or defoliation of Scots pine 

Christmas tree plantings in some eastern and central States and up to 60-percent volume 

reduction in radiata pine (P. radiata) plantings in New Zealand. Serious damage occurs 

only occasionally in some settings in the Great Plains, such as those with highly susceptible seed sources, monocultures, or favorable weather. Retention of two age-classes of 

green needles is considered sufficient for Christmas tree plantations in the eastern and 

Figure 46-3—Cyclaneusma minus fruiting bodies (apothecia) on Scots pine needles: 

(A) Closed apothecia on light tan portion of a needle; and (B) Open apothecia on moist needle 

(James A. Walla, Northern Tree Specialties, used with permission).

Figure 46-2—Cyclaneusma needle cast symptoms on Scots pine needles, showing tan needles with 

brown bars (James A. Walla, Northern Tree Specialties, used with permission).

A B
168 USDA Forest Service RMRS-GTR-335. 2016.

central United States, and that is usually achieved without needle protection in the Great 

Plains. Severe infection results in trees appearing yellow and sparse, thereby reducing 

aesthetic value, growth, and windbreaking ability. Heavily infected young trees often 

have uniform yellowing of second-year needles throughout the tree, not decreasing from 

the base to the top of the trees. The most extensive damage observed in the Great Plains 

was on Scots pine in neglected Christmas tree plantations in South Dakota, and in a 

crowded 11-year-old planting in North Dakota. No damage has been apparent on other 

infected pine species in the Great Plains. C. minus does not typically cause needle cast in 

Europe, but the fungus can still be recovered as an endophyte from some pines.

Management

Control by fungicides is possible, but several applications are needed. In Pennsylvania 

chlorothalonil applied five times (late March, early May, mid-June, early August, early 

October) gave good control. Other fungicides also prevent infection, but their reduced 

longevity on the needles reduces their cost effectiveness. Time of the first application 

should be when buds of common lilac have swollen so about 1/16 inch of green tissue is 

showing next to the bud scales. Two years of protection are needed to restore good needle 

density to sparse trees. If there is a nearby source of infection, annual protection is needed 

in nurseries to prevent distribution of infected seedlings. Such nurseries should work to 

eliminate sources of infection.

Silvicultural practices that increase air movement through plantings, or increase 

distances between susceptible trees, should reduce amount of infection. Improving soil 

fertility may delay symptom development long enough to provide another age-class of 

green needles. There is considerable variation in susceptibility to this fungus among Scots 

pine provenances.

Selected References

Merrill, W.; Wenner, N.G. 1996. Cyclaneusma needlecast and needle retention in Scots 

pine. Plant Disease. 80: 294–298.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.
USDA Forest Service RMRS-GTR-335. 2016. 169

47. Phomopsis Blight of Junipers

Glenn W. Peterson (Riffle and Peterson 1986) (updated by Ned 

Tisserat)

Phomopsis blight is a serious problem in nurseries producing juniper (Juniperus

spp.) seedlings and grafts. It also causes damage to several juniper species in landscape 

plantings.

Hosts and Distribution

Phomopsis blight, caused by the fungus Phomopsis juniperovora, is widespread on 

eastern redcedar (J. virginiana) and Rocky Mountain juniper (J. scopulorum) in the 

eastern United States and the eastern half of the Great Plains, but rare in the more arid, 

western Great Plains. The fungus may also cause damage to creeping (J. horizontalis) 

and savin (J. sabina) junipers, as well as some species of cedar (Cedrus), false-cypress 

(Chamaecyparis), and arborvitae (Thuja).

Symptoms and Signs

P. juniperovora initially infects foliage, then spreads to and sometimes kills stem 

tissues. Newly developing needles are especially susceptible while they are still in the 

yellowish-green stage. After needles become a normal, deep green, they are no longer 

susceptible. Small yellow spots appear on young needles of eastern redcedar and Rocky 

Mountain juniper within three to five days of infection. 

The fungus ramifies within infected needles and rapidly invades and girdles young stems (fig. 47-1). When 

a side shoot is infected, the fungus progresses to the 

main stem, which it may girdle if the stem is less than 

0.4 inches in diameter. The portion of the plant above 

the girdled area then dies.

At first, infected tissues turn light green but then 

rapidly change to the characteristic red-brown color 

of dead shoots, which finally turn ash gray (fig. 47-2). 

Lesions on larger stems frequently develop into cankers, but the stems are not girdled. 

Symptoms of Phomopsis closely resemble those 

of Kabatina blight, and the two diseases may be difficult to distinguish in the field. However, symptoms 

of Phomopsis blight normally appear in spring and 

continue throughout the summer months, whereas 

Kabatina blight symptoms are usually visible by late 

winter.

Disease Cycle

Spores produced in fruiting bodies 

(pycnidia) formed on leaves and stems 

infected the previous year are the most 

important source of inoculum early 

in the growing season. Pycnidia with 

viable spores may develop within three 

to four weeks after seedlings become 

infected, but usually are not well developed until infected tissues have dried 

considerably. Pycnidia are found most 

commonly on tissues that have turned 

ash gray. The pycnidia are embedded at 

first in needles and stems, but partially 

Figure 47-1—Phomopsis blight 

damage (U.S. Forest Service, 

Bugwood.org, used with 

permission).

Figure 47-2—Foliar symptoms of Phomopsis 

juniperovora on eastern redcedar (Bruce Watt, 

University of Maine, Bugwood, used with 

permission).
170 USDA Forest Service RMRS-GTR-335. 2016.

erupt through the epidermis (fig. 47-3). Two types of 

spores (alpha and beta) develop in the same or different 

pycnidia (fig. 47-4) and are extruded in whitish tendrils. 

The alpha spores are clear (hyaline), one-celled, and ellipsoid; contain two oil globules; and commonly are 7.5 to 

10.0 μm × 2.2 to 2.8 μm. The beta spores are hyaline, onecelled, filamentous, slightly curved, and commonly 20.2 to 

26.9 μm × 1.0 μm. The fungus can produce spores for as 

long as two years in dead parts of infected plants. Spores 

are dispersed primarily by rainsplash.

Damage

Damage is most severe in seedling and transplant beds 

of eastern redcedar and Rocky Mountain juniper in nurseries. Total loss of first-year seedlings can occur if control 

measures are not used. Losses are particularly high in 

areas where water tends to stand, and in beds of new seedlings adjacent to infected older seedlings. Survival of even 

lightly blighted nursery stock is very poor, because new 

shoots continue to be infected by spores 

produced on old, infected tissues.

Damage to junipers in landscape plantings is more severe on shrub forms or in 

areas with poor air movement. Trees may 

become unsightly because of numerous 

dead branch tips, but older trees seldom are 

killed because only small-diameter stems 

are girdled. For this reason, Phomopsis 

blight does not cause significant damage in 

natural stands of junipers.

Management

Because susceptible new foliage and 

viable fungal spores are present throughout 

the growing season in juniper seedling beds, protective fungicides need to be applied at 

seven- to 14-day intervals. Avoid poorly drained areas because losses are often greater 

where water tends to stand. If using overhead sprinklers, irrigate seedlings so that water 

on seedlings dries before nightfall. Infected seedlings should be rogued from the nursery 

bed. Junipers or other hosts of this fungus should not be used in nursery windbreaks or 

in landscape plantings on nursery grounds because they may be a source of inoculum 

(spores) for nursery stock. Such trees are more likely to be extensively infected if pruning 

results in development of juvenile foliage.

Phomopsis blight is usually not a serious problem on mature juniper trees in the landscape, but it may cause unsightly damage to shrub or prostrate forms in planting beds and 

require fungicide applications as outlined. In most cases, however, removal of dead tips 

in early spring and during dry weather is sufficient to control this disease. Susceptibility 

to P. juniperovora varies considerably among junipers. Consider choice of plant material 

when planting in disease-prone locations. 

Selected References

Peterson, G.W. 1973. Infection of Juniperus virginiana and J. scopulorum by Phomopsis 

juniperovora. Phytopathology. 63: 246–251.

Peterson, G.W. 1975. Phomopsis blight of junipers. In: Forest nursery diseases in the United 

States. Agric. Handb. 470. Washington, DC: U.S. Department of Agriculture: 76–79.

Peterson, G.W.; Hodges, C.S., Jr. 1982. Phomopsis blight of junipers. Forest Insect and 

Disease Leaflet 154. Washington, DC: U.S. Department of Agriculture, Forest Service. 7 p.

Figure 47-4—Alpha (ellipsoidal) and beta (long) 

spores of Phomopsis species (Lorraine Graney, 

Bartlett Tree Experts, used with permission). 

Figure 47-3—Black fruiting structures (pycnidia) of 

Phomopsis juniperovora

(Robert Anderson, U.S. Forest 

Service).
USDA Forest Service RMRS-GTR-335. 2016. 171

48. Rhizosphaera Needle Cast of Spruce

James A. Walla; revised from Darroll D. Skilling and James A. Walla 

(Riffle and Peterson 1986)

Rhizosphaera needle cast is caused by the fungus Rhizosphaera kalkhoffii. It had 

been the most important needle disease of spruce (Picea spp.) in North America until 

the similar-appearing Stigmina needle cast developed. Needle discoloration caused by 

insects, mites, environmental stress, chemicals, cankers, and normal old-age senescence 

may appear similar to that caused by Rhizosphaera needle cast.

Hosts and Distribution

Hosts include blue (P. pungens), Engelmann (P. engelmannii), Norway (P. abies), 

white (P. glauca, including Black Hills seed sources), and several other spruce species. 

Other conifers, including Douglas-fir (Pseudotsuga menziesii), fir (Abies spp.), and pine 

(Pinus spp.), may also serve as hosts. In North America, Rhizosphaera needle cast has 

been most important in the Great Lakes region and northeastern United States. It occurs 

in the northern Great Plains and causes concern after consecutive years of above-average 

rainfall. It occurs in natural blue and Englemann spruce stands in Arizona and Colorado.

Symptoms and Signs

Faint yellow bands typically develop on recently infected needles in fall or the following spring, four to 11 months after infection. Small dark brown or black spherical fruiting 

bodies (pycnidia; fig. 48-1A) emerge through the stomata the spring after infection. They 

can easily be seen with a hand lens. On severely infected needles, a high proportion 

of stomatal pits contain pycnidia, giving the appearance of continuous fine black lines 

on each side of the needle (fig. 48-2B). White wax plugs from the stomatal pits often 

sit atop the pycnidia. 

The pycnidia contain 

asexual spores (conidia; 

fig. 48-3), which are 

aseptate, oval, colorless 

(hyaline), and small 

(4.5 to 8.6 μm × 2.5 to 

4.6 μm). The needles 

may become fully 

discolored by early summer of the year after 

Figure 48-1—Rhizosphaera kalkhoffii fruiting bodies (pycnidia) 

(A) in stomatal pits (James A. Walla, Northern Tree Specialties, 

used with permission); and (B) showing smooth surface, dark hyphal strands extending from the base, and two fruiting bodies with 

white wax plugs from the underlying stomatal pits (Kasia Kinzer, 

North Dakota State University, used with permission).

A B

Figure 48-2—Rhizosphaera needle cast in spruce trees showing (A) a reduced number of healthy 

needle age-classes, and (B) rows of black fruiting bodies (pycnidia) visible without magnification in 

heavily infected trees (James Walla, Northern Tree Specialties, used with permission) 

A B
172 USDA Forest Service RMRS-GTR-335. 2016.

Figure 48-3—Rhizosphaera 

kalkhoffii (A) pycnidia (dark 

brown structures) and neck 

where spores emerge from 

the pycnidium, and (B) small 

oval spores (stained with 

aniline blue) and hyphal 

strands at the base of a 

pycnidium (James Walla, 

Northern Tree Specialties, 

used with permission).

Figure 48-4—Spruce trees with Rhizosphaera needle cast showing (A) hollow, sparse appearance; 

and (B) wide variation in host resistance, as reflected in amount of defoliation under high disease 

pressure (James A. Walla, Northern Tree Specialties, used with permission).

A B

A B

infection and be cast later that year. The disease typically begins close to the ground in 

the interior crown and progresses upward and outward if conditions are favorable for 

development. Needles of all ages may be infected, sometimes resulting in a mixture of 

symptoms on a given age-class of needles (fig. 48-2A). Trees that have had severe infection for three or more years have a sparse, hollow appearance and may have reduced 

shoot growth (fig. 48-2A) and dead branches in the lower crown (fig. 48-4A).

Blue spruce exhibits a wide range of apparent resistance among trees. Resistance 

levels are expressed as stepwise increases in the number of needle age-classes that are 

green. Under severe disease pressure, the most affected branches on susceptible trees 

will have one age-class of green needles. The next most susceptible trees will have two 

age-classes of green needles, and so on. With severe disease pressure, trees that have four 

or more age-classes of green needles have good resistance to Rhizosphaera needle cast 

(fig. 48-4B).

Stigmina lautii causes a similar needle cast on spruce that can be confused with 

Rhizosphaera needle cast because its sporodochia very closely resemble of R. kalkhoffii. 

Using a 20× hand lens allows differentiation of the two fungi (compare fig. 48-1A with 

fig. 49-2).

Disease Cycle

Conidia are released from mature pycnidia for about two months after budbreak during 

periods of wet weather and are dispersed primarily by rainsplash to cause new infections. 

Symptoms become readily visible the spring of the year after infection. Pycnidia develop 

that spring and produce conidia to complete the disease cycle in one year. The disease 

cycle is longer on more-resistant trees.
USDA Forest Service RMRS-GTR-335. 2016. 173

Damage

Rhizosphaera needle cast can infect and damage trees of all ages, from nursery 

seedlings to mature trees. Blue and Engelmann spruce are the most susceptible species, 

white spruce is intermediate in susceptibility, and Norway spruce is relatively resistant. 

Although some tree mortality has been observed under epidemic conditions, the primary 

damage involves premature needle cast. Branches that have less than three age-classes of 

healthy needles for three to four years will die. Trees with sparse foliage or dead branches 

are unsightly for ornamental purposes, are unmerchantable for high-quality Christmas 

trees, and provide reduced benefits in conservation plantings. In the northern Great 

Plains, consecutive years with greater-than-normal spring rainfall have resulted in serious 

defoliation and branch dieback owing to Rhizosphaera needle cast on blue and sometimes 

white spruce. However, because spruce may be damaged in many ways, it is important to 

realize that presence of a particular pathogen does not mean it is responsible for significant needle discoloration and loss. 

Management

As the fungus can be transported on nursery stock, avoid introducing infected stock 

into plantings. Young spruce planted near older spruce are prone to severe damage even 

when disease is not severe on the older trees. During outbreaks, the disease becomes 

widespread. In areas where this disease occurs sporadically, disease severity fluctuates 

widely with amount of late spring precipitation. Two dry years can reduce disease to 

a minimum, whereas a few wet years can result in severe needle cast. Awareness of 

such fluctuations can help in planning management strategies. If the disease has been a 

problem in the area, consider planting less-susceptible spruce species or other types of 

trees. Promote good air circulation (wide spacing, pruning lower spruce branches and 

overhanging trees, mowing grass and brush). Consider removing trees that are repeatedly 

damaged. Blue spruce plantings should be examined frequently for signs of disease. If 

the trees are dropping their needles prematurely, determine the cause. Early identification helps avoid damage because light infections can be controlled with only one or two 

fungicide sprays. 

Two years of fungicide protection of needles for the first two months after budbreak 

will usually restore moderately infected trees to good foliage quality. Heavily infected 

trees will require more years of treatment. Several fungicides are registered specifically 

for Rhizosphaera needle cast management in the United States and Canada (a, b, respectively, in the following list). Some products containing azoxystrobin, chlorothalonila

, b, copper hydroxidea

, copper oxychloride, copper salts of fatty and resin acids, copper 

sulfate, mancozeba, mono- and di-potassium salts of phosphoric acid, propiconazoleb, 

pyraclostrobin, sulfur, thiophanate-methyl, and trifloxystrobinb are labeled for use on 

spruce and have been found to control some foliar diseases. Chlorothalonil is most commonly recommended, but recent research on efficacy is lacking. In the United States, 

fungicides may be used for disease control if they are registered for the intended target 

site in the state where the application occurs. The products for use in Canada are for commercial or agricultural use.

Selected References

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Walla, J.A. 1990. Variable defoliation levels in Picea pungens by Rhizosphaera 

kalkhoffii. In: Recent research on foliage diseases; conference proceedings; 

May 29–June 2, 1989; Carlisle, PA. Gen. Tech. Rep. WO-56. Washington, DC: U.S. 

Department of Agriculture, Forest Service: 80–82.
174 USDA Forest Service RMRS-GTR-335. 2016.

49. Stigmina Needle Cast of Spruce

James A. Walla and Aaron D. Bergdahl

Stigmina needle cast is caused by the fungus Stigmina lautii. The fungus was first described from native black (Picea mariana) and white (P. glauca) spruce in Manitoba and 

Saskatchewan in 1973. It was first reported causing disease in North Carolina in 2002. 

This disease is often misdiagnosed as Rhizosphaera needle cast.

Hosts and Distribution

Blue (P. pungens) and white (including Black Hills seed sources) spruce are generally 

highly susceptible to Stigmina needle cast, and the disease has been reported on black 

and Norway (P. abies) spruce. Stigmina needle cast was rarely recognized before 2006, 

but it is now known to be widespread east of the Rocky Mountains in the United States 

and Canada.

Symptoms and Signs

One or more faint yellow bands and small patches of tiny round black fruiting bodies 

(sporodochia), which replace the white wax in the stomatal pits, develop in the fall of the 

same year as infection or early the next spring. Either bands or sporodochia may appear 

first. By late summer of the year after infection, individual bands or whole needles may 

be yellow, purple, tan, reddish brown, or 

brown, and sporodochia may occur in more 

than half of the stomatal pits. It appears that 

multiple infections are necessary before 

whole needles become discolored. Wax 

from the stomatal pits may rest atop sporodochia. Most needles are discolored and 

sporodochia mature to produce spores either 

late in the second season, or early in the 

third season after infection (fig. 49-1). With 

dark spores produced mostly horizontal to 

the needle surface, sporodochia appear en 

masse as discontinuous fuzzy black lines 

along the length of each side of a needle, 

especially on green needles (fig. 49-2). All 

needle age-classes are usually susceptible, 

so needles in older age-classes may exhibit 

multiple stages of disease development. 

Microscopic observation of the spores is 

required to confirm Stigmina needle cast. 

Spores of S. lautii are light brown, elongate, 

five- to eight-septate, and 30 to 45 μm × 5.0 

to 5.5 μm when observed with a compound 

microscope (fig. 49-3).

On most heavily infected trees, the two 

youngest age-classes of needles (first- and 

second-year needles) are green or mostly 

green until summer’s end (fig. 49-4). 

Most third-year and any older needles are 

necrotic. Near summer’s end, discoloration 

expands on second-year needles so that 

only first-year needles are bright green. 

Infected needles may drop off shoots as 

soon as the end of the second season after 

infection, but they typically begin dropping near the end of the third summer after 

Figure 49-1—Yellow and brown necrosis and black stippling (fruiting bodies of 

Stigmina lautii) on spruce needles (James A. 

Walla, Northern Tree Specialties, used with 

permission).

Figure 49-2—Fruiting bodies (sporodochia) 

in and over some stomatal pits of a spruce 

needle. Asexual spores (conidia) extend out 

from the sporodochia, resulting in the appearance of fuzzy fruiting bodies, but some 

sporodochia may not have attached conidia. 

Released conidia are often visible lying on the 

needle surface. (Justin Knott, formerly North 

Dakota Department of Agriculture, used with 

permission).
USDA Forest Service RMRS-GTR-335. 2016. 175

infection. Some or all needles 

may remain attached for another 

year or two, and spores continue 

to be produced on them seasonally. A few blue spruce appear to 

have some resistance to Stigmina 

needle cast. That apparent resistance is expressed as a delay in 

infection or development of signs 

and symptoms by one year, so 

there is one additional age-class of 

asymptomatic needles on heavily 

infected trees with this trait.

Disease Cycle

In the northern Great Plains, 

S. lautii has a two-year life cycle. 

Some spores are produced in very 

late fall of the second season after 

infection, but too late to result in 

new infections. The next spring, 

new spores first develop one to 

two weeks before new shoots 

begin growing. Spore production and infection can occur for 

the rest of the growing season 

whenever temperatures are above 

about 50 °F, but the major infection period is the first two months 

after budbreak. On most trees, 

immature sporodochia develop on 

needles by late spring about one 

year after infection and produce 

spores six to 12 months after 

formation, continuing the disease 

cycle.

Damage

Damage from Stigmina needle 

cast results from the presence of 

discolored needles and the loss 

of functioning needles. Stigmina 

needle cast is proving to be 

more aggressive and to have 

longer-lasting epidemics than 

other spruce needle diseases. 

Thinning of foliage commonly 

occurs over the lower two-thirds 

of the crown of open-grown trees 

(fig. 49-5), and a greater portion 

of tree crowns may be affected 

Figure 49-3—Squash mount of Stigmina lautii fruiting 

body (sporodochium), with brown septate spores (conidia; 

Kasia Kinzer, North Dakota State University, used with 

permission).

Figure 49-4—Spruce shoots with needles showing 

symptoms of Stigmina needle cast. Second-year needles 

have advanced symptoms, and third-year needles have 

been cast (late August; James A. Walla, Northern Tree 

Specialties, used with permission).

Figure 49-5—Discolored needles and sparse foliage 

caused by Stigmina needle cast (James A. Walla, Northern 

Tree Specialties, used with permission).
176 USDA Forest Service RMRS-GTR-335. 2016.

in dense stands. In North Dakota, the impact of Stigmina needle cast increased each year 

from 2006 to 2012, through periods with above-average rainfall and periods with belowaverage rainfall, to a level well beyond that associated with other spruce needle diseases.

Management

Two fungicide management approaches are available. In the first approach, needle 

protection for two months after budbreak, repeated every year, will reduce disease so 

branches retain three or more age-classes of green needles. This approach would be useful for landscape, Christmas, and other trees where loss of the oldest classes of needles is 

acceptable. In the second approach, needle protection for the entire infection period (usually mid-May to mid-October in North Dakota) every year for four or five years, is meant 

to prevent infections until previously infected needles drop from the tree. This approach 

would be necessary for nursery stock where certification requires that plants be diseasefree and for those that prefer to maximize the aesthetic value of a tree. 

No fungicides are labeled specifically for Stigmina needle cast in the United States. 

However, fungicides may be used for disease control in the United States if they are 

registered for the intended target site in the state where the application occurs. For needle 

diseases on spruce, these fungicides include some products containing azoxystrobin, 

chlorothalonil, copper hydroxide, copper salts of fatty and resin acids, copper sulfate, 

mancozeb, mono- and di-potassium salts of phosphoric acid, pyraclostrobin, sulfur, and 

thiophanate-methyl. In Canada, chlorothalonil, propiconazole, and trifloxystrobin are labeled for use by commercial applicators specifically for management of Stigmina needle 

cast in Christmas tree nurseries, and copper-oxychloride is similarly labeled except it is 

registered for management of needle cast diseases in general. In North Dakota, experimental application of chlorothalonil under the first approach (above) was highly effective 

during the protection period, and experimental application of copper sulfate under the 

second approach (above) was moderately effective.

Because potential fungicide management options will be long-term or intensive, 

determine the value of a spruce tree to decide whether disease management is justified. In 

areas where Stigmina needle cast is epidemic, refraining from planting susceptible spruce 

and removal of severely damaged spruce are justified. Stigmina needle cast has not been 

found on a Norway spruce cultivar (‘Royal Splendor’) in North Dakota during multiple 

inspections since 2006, whereas adjacent blue spruce trees were severely diseased. This 

apparent resistance has not been evaluated in replicated trials, however. Cultural methods 

may reduce Stigmina needle cast, but their efficacy has not been established.

Selected References

Sutton, B. C. 1973. Hyphomycetes from Manitoba and Saskatchewan, Canada. 

Mycological Papers. 132: 1–143.

Walla, J.A.; Kinzer, K. 2006. Stigmina lautii discovered on spruce in North Dakota. Tree 

Talk. 2(5): 13–18. http://www.ag.ndsu.edu/trees/whatnew/Tree_Talk_Dec2006-2.pdf. 

(March 6, 2015).
USDA Forest Service RMRS-GTR-335. 2016. 177

50. Canker Diseases of Juniper and Oriental 

Arborvitae

Ned Tisserat

Rocky Mountain juniper (Juniperus scopulorum) and oriental arborvitae (Platycladus 

orientalis) may be seriously damaged by canker diseases, especially in the southern Great 

Plains. 

Hosts and Distribution

Two canker diseases cause dieback and mortality of juniper (Juniperus spp.) 

and oriental arborvitae in the Great Plains. Botryosphaeria canker, caused by the 

fungus Botryosphaeria stevensii, is a serious disease of Rocky Mountain juniper in 

the south-central and southeastern Great Plains. The disease also occurs on eastern 

redcedar (J. virginiana) and savin juniper (J. sabina). The disease has also been 

reported on savin juniper in Pennsylvania and is likely widespread on junipers in the 

eastern United States. A second fungus, Seiridium unicorne (teleomorph Lepteutypa 

cupressi), primarily causes cankers on winter-damaged or drought-stressed oriental 

arborvitae, and occasionally juniper species, primarily from Kansas south into 

Texas. The fungus may occasionally cause cankers on other tree species in the cypress (Cupressaceae) family.

Symptoms and Signs

Junipers affected with Botryosphaeria canker (fig 50-1) develop elongated, flattened, often resinous cankers (figs. 50-2). These cankers may occur anywhere on 

woody stems, but are commonly located near branch crotches in the interior portion 

of the tree crown. Cankers are sometimes difficult to see, and it is often necessary to 

cut the dead branch off and carefully scrape away the outer bark to expose the chocolate-brown dead tissue in the canker in the living bark (phloem). The surrounding 

healthy phloem or inner bark is pearl white. Small black fruiting bodies (pycnidia) 

of the fungus also develop in the canker, but may be partially hidden by thin pieces 

of dead bark. Pycnidia produce spores (conidia; 25 to 31 μm × 10 to13 μm) that are 

at first single-celled and clear (hyaline; fig. 50-3), but when mature become dark 

brown and often two-celled.

Symptoms of Seiridium canker are similar to Botryosphaeria canker, and microscopic examination of the fungal spores may be required to differentiate them. 

Figure 50-1—Branch dieback on (A) Rocky Mountain juniper caused by Botryosphaeria stevensii, and (B) oriental 

arborvitae caused by Seiridium unicorne (Ned Tisserat, 

Colorado State University, used with permission).

A B
178 USDA Forest Service RMRS-GTR-335. 2016.

Figure 50-3—Asexual spores (conidia) of (A) Botryosphaeria stevensii, and (B) Seiridium unicorne

(Ned Tisserat, Colorado State University, used with permission).

A B

Figure 50-2—Flattened branch canker on (A) Rocky Mountain juniper caused by 

Botryosphaeria stevensii, and (B) oriental arborvitae caused by Seiridium unicorne (Ned 

Tisserat, Colorado State University, used with permission).

A

B
USDA Forest Service RMRS-GTR-335. 2016. 179

Seiridium unicorne causes elongated, flattened cankers to form on small branches 

and main stems. On junipers, these cankers may be larger and more diffuse than 

those caused by B. stevensii. Bleeding or resin formation in cankers, especially on 

oriental arborvitae, is common (fig 50-2B). Multiple coalescing branch and stem 

cankers may cause branch dieback or, in some cases, tree death. In the cankers the 

fungus forms black pycnidia, which produce brown-pigmented, multiseptate conidia 

with appendages (fig. 50-3B). 

Disease Cycle

Spores of B. stevensii are dispersed relatively short distances by rainsplash. Peak 

spore release occurs in late May and June, suggesting this is a major period of infection. 

Spores of S. unicorne are similarly released during rain. Both fungi likely infect through 

bark wounds. Cankers develop over several months and typically girdle small branches 

within a year, causing death of the branch distal to the canker. Dead branches scattered 

throughout the crown may result. Infection by S. unicorne is greatly exacerbated in trees 

suffering from winter damage or drought stress, whereas B. stevensii will cause cankers 

even on vigorously growing trees. 

Damage

Occasionally, B. stevensii stem cankers will cause girdling and rapid death of the top 

one-third to one-half of the tree crown. More commonly, the disease causes death of 

individual branches throughout the crown and a gradual tree decline. This fungus can 

kill large vigorous Rocky Mountain junipers in both windbreak and ornamental settings. 

Mortality in windbreak plantings is especially damaging because it may create large gaps. 

Botryosphaeria canker should not be confused with Kabatina or Phomopsis blight, both 

of which affect only foliage and succulent branch tips. Seiridium canker may also kill 

oriental arborvitaes, but more commonly results in scattered branch dieback. 

Management

Rocky Mountain juniper is very susceptible to Botryosphaeria canker and should not 

be planted in locations where the disease is common. Savin juniper also is susceptible, 

but eastern redcedar and Chinese juniper are more resistant. Fungicides are not currently 

labeled for control of Botryosphaeria canker. Remove cankers on diseased trees in winter 

or late spring. Do not prune or shear cankered junipers in May or June, when spores 

are released. Pruning and sanitation may not completely suppress canker development. 

Seiridium canker tends to be associated with trees that are suffering from winter damage, 

drought, or other environmental stresses and thus less able to defend against the fungus. 

Canker development can be suppressed by planting winter-hardy selections of oriental 

arborvitae and protecting trees from winter desiccation. Cankered branches should be 

pruned from trees and destroyed.

Selected References

Stanosz, G.R.; Moorman, G.W. 1997. Branch dieback of savin juniper in Pennsylvania 

caused by Diplodia mutila. Plant Disease. 81: 111. 

Tisserat, N.A.; Nus, A.; Barnes, L.W. 1991. A canker disease of the Cupressaceae in 

Kansas and Texas caused by Seiridium unicorne. Plant Disease. 75: 138–140.

Tisserat, N.A.; Pair, J.C. 1997. Susceptibility of selected juniper cultivars to cedar-apple 

rust, Kabatina tip blight, Cercospora needle blight and Botryosphaeria canker. Journal 

of Environmental Horticulture. 15: 160–163. 

Tisserat, N.A.; Rossman, A.Y.; Nus, A. 1988. A canker disease of Rocky Mountain 

juniper caused by Botryosphaeria stevensii. Plant Disease. 72: 699–701. 
180 USDA Forest Service RMRS-GTR-335. 2016.

51. Diplodia Shoot Blight and Canker Disease

James T. Blodgett and Glen R. Stanosz; revised from Glenn W. 

Peterson and David W. Johnson (Riffle and Peterson 1986)

The fungal pathogen, Diplodia sapinea (D. pinea, Sphaeropsis sapinea), causes shoot 

blights and cankers in pines (Pinus spp.) and some other conifers in much of the United 

States. Damage can be severe in landscapes, windbreaks, parks, and forests. A closely 

related pathogen, D. scrobiculata, occurs in the United States and may be present in the 

Great Plains although it has not been reported.

Hosts and Distribution

Diplodia shoot blight and canker disease is a common and sometimes serious problem 

in the Great Plains. Pines are affected from seedling stage to maturity. The disease affects trees in nurseries, Christmas tree and ornamental plantings, plantations, and natural 

stands. Exotic and native two- and three-needle pines are commonly damaged in the 

United States. D. sapinea is widely distributed in the Great Plains and Diplodia shoot 

blight and canker disease can be locally severe, affecting Austrian (P. nigra), ponderosa 

(P. ponderosa), Scots (P. sylvestris), red (P. resinosa), and mugo (P. mugo) pines. Austrian 

pine, which has been widely used in landscape, windbreak, and park plantings since the 

early 1900s, is especially susceptible.

Symptoms and Signs

Symptoms of Diplodia shoot blight on new shoots first occur during late May to late 

June. The first indications of infection are often resin droplets or one to a few short dead 

needles on emerging shoots, or a combination thereof. The most conspicuous symptom of 

Diplodia shoot blight is death of new shoots containing short, light brown, often wilting 

needles (fig. 51-1) that fade to gray and remain attached to stems. Both needles and stems 

can become stunted or crooked. D. sapinea commonly kills entire new shoots by mid- to 

late summer. Cankers, often resinous, develop in branches and stems with discolored 

brown phloem and cambium. Underlying xylem is often stained blue.

Symptomatic, new shoots may 

be scattered throughout crowns, 

although damage is often first evident in lower crowns. Symptom 

severity can vary considerably 

among major branches. In some 

cases, damage may be confined 

to new shoots (fig. 51-2A), 

particularly on trees affected for 

the first time. After two or three 

successive years of damage, extensive branch and top mortality 

can occur (fig. 51-2B). Repeated 

outbreaks of Diplodia shoot 

blight and canker disease result in 

reduced growth, deformed trees, 

and tree mortality.

Small black asexual fruiting 

bodies (pycnidia) are easily 

observed with a 10× hand lens. 

However, microscopic examination of the asexual spores 

(conidia; fig. 51-3) or DNA 

sequence-based diagnostics are 

necessary for identification. 

Pycnidia are often numerous 

Figure 51-1—Typical symptom of Diplodia shoot blight: 

a dead, crooked new shoot with light brown needles that 

remain attached (James T. Blodgett, U.S. Forest Service).
USDA Forest Service RMRS-GTR-335. 2016. 181

Figure 51-2—Mortality caused by Diplodia shoot blight in (A) shoots, and (B) top/branches of 

ponderosa pine (James T. Blodgett, U.S. Forest Service).

at the base of short, ash-gray 

needles killed the previous year 

(fig. 51-4). Pycnidia can develop 

in stems and may be numerous 

in scales of mature seed cones 

(fig. 51-5). To assist in diagnosis, 

needle and stem samples may be 

incubated in plastic bags with 

a piece of damp paper towel. 

Pycnidia with conidia may develop within two to three weeks.

Disease Cycle

D. sapinea survives as 

mycelium or pycnidia in killed 

needles, bark of young shoots, 

and scales of mature seed cones. 

Numerous conidia are dispersed 

during rainfall, especially in 

spring, but conidial dissemination can also occur throughout the year. Conidia are spread 

by rainsplash, wind-driven rain, or insects.

Needles and young elongating stems are most susceptible to D. sapinea infection in 

spring from budbreak until around mid-June. Conidia germinate and germ tubes can 

directly penetrate nonwounded, immature needle and stem tissues. The pathogen also can 

infect both current-year shoots and older tissues through wounds. Rain or high humidity 

coupled with cool to moderate temperatures favor spore germination, germ tube growth, 

and infection. If rain is sparse when new shoots are highly susceptible, disease incidence 

is usually low. After infection, new shoots are often killed quickly. Pycnidia may be 

produced within two weeks of shoot infection in spring, or throughout the summer in wet 

years.

Seed cones bearing pycnidia are often present on pines that otherwise appear healthy. 

These cones can be a source of large numbers of conidia. The conidia can germinate to 

initiate new infections on lower branches of the tree, nearby trees, and seedlings. Debris 

left onsite after harvesting also may serve as a source of conidia for several years.

This pathogen can persist on or in asymptomatic stems and branches of pines. Water 

stress has been demonstrated to act as a trigger, allowing D. sapinea to proliferate and 

A B

Figure 51-3—Spores (conidia) of Diplodia sapinea with 

light young conidia and dark brown older conidia; approximate size 34 to 39 μm × 12 to 13 μm (James T. Blodgett, 

U.S. Forest Service). 
182 USDA Forest Service RMRS-GTR-335. 2016.

Figure 51-5—Second-year seed cone scales with (A) fruiting bodies 

(pycnidia) of Diplodia sapinea, and (B) uninfected scales (James T. 

Blodgett, U.S. Forest Service).

Figure 51-4—Fruiting bodies (pycnidia) of Diplodia sapinea at the base 

of pine needle with fascicle sheath removed (James T. Blodgett, U.S. 

Forest Service). 

B

A

cause disease in these trees. This ability to act as a latent pathogen could explain the rapid 

mortality observed after outplanting or following hail or other wound events.

Damage

Diplodia sapinea can cause shoot blight, canker, crown wilt, collar rot, and root 

disease. Although shoot blight is the most common type of damage, branch death, top 

dieback, and tree mortality can occur. Diplodia shoot blight and canker disease can 

predispose trees to other diseases and attack by insects, including the pine engraver (Ips

spp.).

Increased damage from Diplodia shoot blight and canker disease has been associated 

with water deficits, competing vegetation, areas with high soil nitrogen, and offsite plantings. This disease often severely affects trees wounded by hail, snow, pruning, shearing, 

insects, or other damage. In the Great Plains, hail events are frequently associated with 

outbreaks of the disease and symptoms develop in one to two weeks. Wounded tissues 

remain vulnerable to D. sapinea infection for several days.

Although pines of all ages are susceptible to D. sapinea, damage is more severe in 

older plantings. In the Great Plains, windbreaks that are 20 to 22 years old have few 
USDA Forest Service RMRS-GTR-335. 2016. 183

diseased pines. However, incidence and damage from Diplodia shoot blight and canker 

disease increase as trees approach 30 years, and damage is often severe in older pines 

with abundant mature seed cones.

Management

Reducing water stress, avoiding wounds, and maintaining tree vigor are often the best 

options to manage Diplodia shoot blight and canker disease. Methods to reduce water 

stress include selecting species suited to the site, proper planting, managing competing 

vegetation, and stand thinning. In developed sites, watering during droughts is recommended. Avoid pruning or shearing in Christmas tree or other pine plantings during 

rain events. Conduct pruning during the dormant season, and disinfect pruning tools. 

Seedlings, including those in nursery beds, and small trees become infected when they 

are located close to diseased or cone-bearing pines. Remove cone-bearing trees or plant 

nonhost trees in the area. Nitrogen fertilization of pines at levels recommended for ornamental and shade trees can result in increased disease incidence and severity, and should 

be avoided.

Selecting nonhost species or more-resistant pine species is an option for long-term 

management. Studies have shown that pines differ in susceptibility, but most two- and 

three-needle pines are susceptible to this disease. Austrian pine is extensively damaged by 

Diplodia shoot blight and canker disease. Although ponderosa pine is susceptible to this 

disease, it is much more tolerant than Austrian, lodgepole, or red pine in the Great Plains. 

Ponderosa, Scots, and jack (P. banksiana) pine do not differ markedly in susceptibility to 

Diplodia shoot blight and canker disease. Scots pine is often recommended for landscape 

plantings, because it frequently experiences less damage from this disease than Austrian 

pine, and ponderosa pine planted offsite.

Infection of new shoots can be reduced significantly by applying an appropriate 

registered fungicide at the time when shoots are highly susceptible, which is usually 

between late April and mid-May, from budbreak to the end of shoot expansion. Fungicide 

applications will protect new shoots, but will not prevent infection of cones or eliminate 

the pathogen in colonized tissues. Fungicide treatments are usually applied to nursery 

seedlings, Christmas trees, or individual high-value landscape trees. Consult your state or 

local extension agent regarding appropriate registered fungicides.

Selected References

Blodgett, J.T.; Stanosz, G.R.; Kruger, E.L. 1997. Sphaeropsis sapinea and water stress in 

a red pine plantation in central Wisconsin. Phytopathology. 87: 429–434.

Flowers, J.; Nuckles, E.; Hartman, J.; Vaillancourt, L. 2001. Latent infection of Austrian 

and Scots pine tissues by Sphaeropsis sapinea. Plant Disease. 85: 1107–1112.

Peterson, G.W. 1978. Effective and economical methods for controlling diplodia tip 

blight. American Nurseryman. 147: 13, 66, 70, 72. 

Peterson, G.W. 1981. Diplodia blight of pines. Forest Insect and Disease Leaflet 161. 

Washington, DC: U.S. Department of Agriculture, Forest Service. 7 p.
184 USDA Forest Service RMRS-GTR-335. 2016.

52. Gymnosporangium Rusts

Simeon Wright, James A. Walla, and William R. Jacobi; revised from 

James A. Walla and Jerry W. Riffle and from Kenneth E. Conway and 

Mark W. Andrews (Riffle and Peterson 1986)

Many Gymnosporangium species occur in the Great Plains. Because it is difficult to 

differentiate them based on symptoms, diagnosis of species is often imprecise. Species 

identification is primarily based on size and shape of the various spores, the morphology 

of fruiting bodies on rosaceous hosts (aecia) and fruiting bodies on Juniperus hosts (telia), and on host symptoms. There is much overlap in these traits among species, but each 

species has a unique combination of the traits. Molecular diagnostics using telial horns or 

masses of aeciospores offer increasingly useful diagnostic capability. It may not be necessary to identify the rust species unless management involves removing an alternate host 

or planting resistant varieties.

The use of common names for diseases caused by these fungi does not always exactly 

fit the hosts being diagnosed. For example, the name cedar apple rust refers specifically 

to the disease caused by G. juniperi-virginianae on junipers and apples (Malus spp.). In 

this case, “cedar” is derived from the partial common name of one of the plants that is 

infected by the fungus, eastern redcedar (J. virginiana), which is a juniper. However, this 

fungus also infects other hosts.

Hosts and Distribution

Several host-specific Gymnosporangium rust species occur in the Great Plains 

(table 52-1). These species alternately infect two unrelated hosts to complete their life 

cycle. The aecial stage almost always occurs on rosaceous species and the telial stage occurs on Juniperus species. Rust species of particular significance to the Great Plains are 

individually discussed. 

Symptoms and Signs

Symptoms vary with the rust species (table 52-1). Symptoms on rosaceous hosts 

include yellow, orange, or red lesions on leaves (fig. 52-1), petioles, fruit (fig. 52-2), or 

new twigs (fig. 52-3) and rough, elongate cankers or gall-like swellings on leaf veins, 

petioles, twigs, and branches. Fruit can be distorted, and premature leaf drop and dieback 

can occur. The yellow to black pimplelike sexual fruiting bodies (spermagonia) (fig. 

52-4) and later tubelike aecia develop in rosaceous host lesions. Symptoms on Juniperus

hosts include small lesions on needles, globular branch galls (fig. 52-5), spindle-shaped 

stem swellings (fig. 52-6), witches’-brooms (fig. 52-7), and branch dieback. Gelatinous, 

orange, fingerlike telial horns (fig. 52-8) or cushionlike telial masses grow and swell from 

infections on the Juniperus hosts in spring.

Disease Cycle

Spring rains are absorbed by each lesion, gall, swelling, or witches’-broom on junipers, causing telial horns or cushionlike masses to swell. Swelling may begin as early as 

prebloom or as late as full leafout of the rosaceous hosts, depending on the rust species, 

its hosts, and the environmental setting. Teliospores in the swollen telial structures germinate to produce basidia that bear basidiospores, which are carried by wind or insects 

to nearby rosaceous hosts. Telial horns or cushions may may swell and dry down several 

several times with repeated wetting, allowing production of basidiospores for eight to 10 

weeks. After infection of susceptible host tissue by basidiospores, spermagonia develop 

in yellow to orange lesions, followed by production of aecia. Aeciospores are blown by 

wind to junipers from late spring to early fall, depending on the species of rust fungus. 

After infection of the juniper, galls or other structures form during the next one to two 

years. New telial horns or masses appear the next spring and the cycle repeats. Some 

galls, swellings, or witches’-brooms produce telial horns or masses annually thereafter.
USDA Forest Service RMRS-GTR-335. 2016. 185

Table 52-1—Some Gymnosporangium species in the Great Plains

Rust species Distribution Symptoms on junipers Major aecial hostsa

G. bethelii Widespread in Rough, irregular, Hawthorn, crabapple, 

west spindle-shaped galls apple

G. clavariiforme Widespread Stem swellings, Amelanchier spp.

 witches’-brooms

G. clavipes Widespread Spindle-shaped stem Many Rosaceae

 swellings

G. connersii North Branch galls Hawthorn

G. corniculans North Branch galls Amelanchier spp.

G. exiguum South Foliar lesions Hawthorn

G. globosum Widespread Branch galls Hawthorn

G. gracile Texas Stem swellings, Amelanchier spp.

 witches’-brooms

G. juniperi- Widespread Branch galls Apple, crabapple

virginianae

G. nelsonii Western and Branch gall or no Amelanchier spp.

north symptoms

G. nidus-avis Widespread Stem swellings, Amelanchier spp.

 witches’-brooms

G. trachysorum South Branch galls Hawthorn 

a Other Rosaceae may be aecial hosts for some of these rusts.

Figure 52-1—Lesions and fruiting bodies (spermogonia, aecia) on hawthorn leaves (David 

Leatherman, Colorado State Forest Service, 

used with permission).

Figure 52-2—Fruiting bodies (aecia) on hawthorn 

leaves and fruit (Harold Larsen, Colorado State 

University, used with permission).
186 USDA Forest Service RMRS-GTR-335. 2016.

Damage

Damage is not usually life threatening on rosaceous hosts, although severe infection 

reduces yield, fruit quality, vigor, and aesthetic value. Apples and Amelanchier species 

(serviceberry, juneberry, Saskatoon berry) are particularly susceptible to these effects.

Damage to junipers is usually slight with reduced vigor and aesthetic value. Rust 

fungi that cause stem swellings or galls may kill branches above the infected points 

when the swellings or galls die. If numerous large galls occur on a juniper, branches 

can be permanently bent downward. Rusts that cause witches’-brooms can deform trees 

and serve as nutrient sinks to reduce vigor in the rest of the plant. If broadleaf hosts are 

nearby and environmental conditions favor infection, spectacular numbers of galls or 

witches’-brooms can form on junipers. Susceptible groundcover junipers have been killed 

by branch-girdling galls.

The following rusts are 

common in one or more 

areas of the Great Plains. 

Information included in 

the descriptions provides 

an indication of the range 

of symptoms that may 

be encountered among 

Gymnosporangium rusts and 

the types of field observations 

that may be used to provide 

rough diagnosis.

Cedar Apple Rust

G. juniperi-virginianae is widespread east of the Rocky Mountains and has been 

reported in some western Great Plains States. The primary aecial hosts are apple and crabapple, where yellow lesions appear on foliage beginning in late spring or early summer. 

Mature lesion color varies from yellow to orange and red. Lesions may also appear on 

fruit and, rarely, on twigs. Short aecia (1 to 2 mm long) develop in midsummer, open by 

aecial walls flaring back from the center, and release aeciospores until fall. Aeciospores 

infect leaves of eastern redcedar, Rocky Mountain juniper (J. scopulorum), and various 

ornamental junipers. Over the next year, brown galls measuring 0.25 to 2.0 inches in 

diameter develop on twigs of the juniper host, followed by long, pointed telial horns 

on galls the following spring. The galls die after one season of sporulation. Resistance 

levels of many apple, crabapple, and 

juniper cultivars to this rust are commonly available in extension bulletins 

and fruit production manuals. The rust 

has multiple races, however, and cultivar 

resistance can change if different races 

become prevalent in a given area.

American Hawthorn Rust

G. globosum is widespread in the 

eastern Great Plains, and has been reported in some prairie provinces and western 

Great Plains states. The primary aecial 

hosts are apple, crabapple, and hawthorn 

(Crataegus spp.), although several 

other rosaceous species are susceptible. 

Infections of aecial hosts may occur on leaves, fruit, petioles, and twigs, but fruit infection is rare. Mature lesions appear in late spring and vary in color from yellow to orange 

and red. In contrast with cedar apple rust, long (3 to 5 mm) tubelike aecia extend from 

lesions in midsummer, open by aecial walls tearing irregularly, and release aeciospores 

until fall. Aeciospores infect leaves of eastern redcedar, Rocky Mountain juniper, and 

Figure 52-3—Fruiting bodies (aecia) in leaf pustules and stem 

swellings on serviceberry (James A. Walla, Northern Tree 

Specialties, used with permission).

Figure 52-4—Fruiting bodies (spermagonia) on 

hawthorn leaves, stems, and fruit (Harold Larsen, 

Colorado State University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 187

various ornamental junipers. Over the next year, brown galls (0.1 to 0.6 inch diameter) 

develop on twigs of the juniper host, followed by short telial horns the following spring. 

In contrast with cedar apple rust, the galls are smaller, can produce spores for multiple 

years, and frequently do not kill twigs.

Quince Rust

G. clavipes is widespread in the eastern Great Plains and has been reported in the 

prairie provinces and some western Great Plains States. Eleven genera of rosaceous 

plants are aecial hosts: Amelanchier, Aronia (chokeberry), Chaenomeles (flowering 

quince), Cotoneaster (cotoneaster), Crataegus, Cydonia (common quince), Malus, 

Mespilus (medlar), Photinia (photinia), Pyrus (pear), and Sorbus (mountain-ash). 

Symptoms vary but frequently include fruit distortion and swelling of leaf midveins, 

petioles, and new shoots. Thorns and young branches are also susceptible and may develop elongate, spindle-shaped perennial cankers. White tubular aecia, 2 to 3 mm long, 

develop during the summer, becoming torn at the tip or splitting along the side to release 

aeciospores until fall. Aeciospores infect leaves and soft stems of eastern redcedar, Rocky 

Mountain juniper, and certain other juniper species. Slight swelling develops around the 

obscure young infections. It is unknown whether cushionlike telia are produced during 

the first or second spring and early summer after infection. Most of the infections die 

after sporulation, but those 

that survive develop into 

elongate, swollen (spindleshaped) perennial cankers 

on twigs and branches. 

Perennial cankers often die 

within four to six years. 

Those that survive can 

cause stem swellings 8 

to 10 inches in diameter. 

Severe infection of junipers can reduce winter 

hardiness.

Figure 52-5—Globular branch galls on juniper (left: William R. Jacobi, Colorado State University, 

used with permission; right: Simeon Wright, Missouri Department of Conservation, used with 

permission).

Figure 52-6—Spindle-shaped stem swelling with cushionlike telia 

on juniper (Simeon Wright, Missouri Department of Conservation, 

used with permission).
188 USDA Forest Service RMRS-GTR-335. 2016.

Juniper Broom Rust

G. nidus-avis is widespread in North America. Several species of serviceberry, 

including Amelanchier alnifolia and A. stolonifera, are susceptible, as are common 

quince and some flowering quince and apple species. Aecia 2 to 5 mm long develop in 

mid-to-late summer on leaves, petioles, new shoots, and fruit and tear irregularly when 

mature. Aeciospores infect leaves and green shoots of various junipers, including Rocky 

Mountain juniper, eastern redcedar, creeping juniper (J. horizontalis), and Chinese juniper (J. chinensis). On the juniper host, small witches’-brooms may develop throughout 

the crown. Some of them may become perennial to form one to several witches’-brooms 

more than 2 ft in diameter. In perennial 

witches’-brooms, the shoots are dwarfed 

and highly branched, and there is often 

a reversion of the needles to the juvenile 

form. Sometimes portions of the subtending branch are swollen. Infection of stems 

may lead to spindle-shaped stem swellings 

without witches’-brooms. Small orange 

gelatinous telial masses appear on leaves, 

shoots, and stems throughout the witches’-

brooms and stem swellings in spring to early 

summer.

Japanese Apple Rust

G. yamadae is a damaging exotic rust 

from Asia that was first observed in the 

United States in 2004. As of 2013, it was 

limited to nine eastern states and one 

province. The primary aecial hosts are 

apple species, and the primary telial host is 

Chinese juniper. On apple leaves, spots are 

bright red with a pale cream to white center. 

Infection of fruit is rare. Severe infection 

occurs on susceptible apple varieties, but 

susceptibility of apple varieties resistant to 

cedar apple rust is unknown. G. yamadae

is morphologically similar to G. globosum, 

but the aecial side walls and telial horn 

lengths are differentials. Aecia on the lower 

apple leaf surface are 3 to 7 mm long. On 

junipers, globoid swellings or small galls are 

produced on branches. Telial masses 0.2 to 

0.3 inches long protrude from swellings like 

rubbery shelves. The pathogen dies after one 

season of sporulation.

Juniper Knot Gall Rust 

G. bethelii is very common on the 

plains of eastern Colorado. This rust forms 

reddish-brown, irregular, perennial galls 0.1 

to 0.6 inches in diameter on several juniper 

species. Telia are chestnut brown, wedge 

shaped, and 0.1 to 0.2 inches high. The alternate hosts are primarily hawthorn, apple, 

crabapple, and others. The aecia produce 

long hair-like tubes on the underside of 

leaves.

Figure 52-7—Witches’-brooms in juniper host, 

showing wet and dry telia (top, middle: James 

A. Walla, Northern Tree Specialties, used with 

permission; bottom: Harold Larsen, Colorado 

State University, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 189

Management

Rust management is rarely required on junipers with galls, but large witches’-brooms 

damage junipers and should be removed. A combination of practices may effectively 

manage these diseases on rosaceous hosts.

Damage can be reduced by not planting susceptible varieties near alternate hosts. 

Varieties may be resistant to some rust species and not to others. Susceptibility of apple, 

crabapple, hawthorn, and juniper varieties and species has been determined for some of 

the most common rusts. When only a few branches are affected, pruning galls or other 

telia-producing structures from Juniperus hosts may reduce infection on alternate hosts. 

Eradication of the Juniperus host from the vicinity may reduce new infections on the 

rosaceous host because basidiospores are fragile. A two mile separation may be required 

to minimize rust infections, but a half mile separation will substantially reduce infection.

Fungicides are registered for both rosaceous and Juniperus hosts. Application timing 

and coverage are critical. Make fungicide applications to protect rosaceous hosts according to the fungicide label. The prescribed timing is generally as flower buds expand 

until telial structures on alternate Juniperus hosts become inactive in late spring or early 

summer (usually four to six weeks). Fungicide applications to protect Juniperus hosts are 

rarely necessary and are challenging to time appropriately. Repeated fungicide applications according to the label from early July through August may help prevent infection. 

Selected References

Kern, F.D. 1973. A revised taxonomic account of Gymnosporangium. University Park, 

PA: Pennsylvania State University Press. 134 p.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.

Yun, H.Y. 2010. Japanese apple rust. Invasive fungi. Beltsville, MD: U.S. Department 

of Agriculture, Agricultural Research Service. http://nt.ars-grin.gov/sbmlweb/fungi/

index.cfm. (July 8, 2014).

Figure 52-8—Gelatinous telial horns 

swelling from juniper galls in spring 

(top: James A. Walla, Northern Tree 

Specialties, used with permission; 

bottom: James T. Blodgett, USDA 

Forest Service).
190 USDA Forest Service RMRS-GTR-335. 2016.

53. Valsa (Cytospora) Canker of Spruce

James A. Walla and Aaron D. Bergdahl; revised from James A. Walla 

and Frederick J. Crowe (Riffle and Peterson 1986)

Valsa canker is caused by the fungus Valsa kunzei (syn. Leucostoma kunzei). The 

imperfect stage is Cytospora kunzei (syn. Leucocytospora kunzei).

Hosts and Distribution

Many species of spruce (Picea spp.) are susceptible to Valsa canker, including blue 

(also called Colorado blue; P. pungens), Engelmann (P. engelmannii), Norway (P. abies), 

and white (P. glauca; including Black Hills seed sources) spruce. Valsa canker may also 

occur on Douglas-fir (Pseudotsuga menziesii), larch (Larix spp.), pine (Pinus spp.), and 

other conifers. In North America, it has been reported throughout the Upper Midwest and 

northeastern United States and adjacent Canada, and in the mountains of Colorado. It occurs in the Prairie Provinces, and the central and northern Great Plains states.

Symptoms and Signs

Branch dieback often occurs first on individual lower branches and then sporadically 

on other branches laterally and upward in the tree (fig. 53-1). In the northern Great 

Plains, the first outward symptoms usually occur in spring, when needles of branches 

girdled since late summer of the previous year begin turning grayish green (fig. 53-1A) 

and then light purple to brown (fig. 53-1B) as they dry out with warming temperatures. 

Branch girdling sometimes occurs from late spring to midsummer, resulting in development of the first outward symptoms during summer. One or more months after becoming 

discolored, needles drop from girdled branches. The youngest blue spruce twigs on dead 

branches typically remain orange for about one year after needles drop, and then become 

gray. Resin exudes from cankered branches and may drip onto noncankered branches. 

Figure 53-1—Colorado blue spruce trees with Valsa canker. (A) Cankered branch with fading 

(gray-green) needles (arrow), early August; (B) Same branch in late September when needles have 

turned brown and started falling (arrow). Twigs remain orange for several months after needles have 

dropped (James A. Walla, Northern Tree Specialties, used with permission); (C) Serious damage by 

Valsa canker on blue spruce, showing recently dead branch with brown needles, branch that died the 

previous year with bleached gray twigs (arrow), and open areas where branches have been removed 

(Aaron D. Bergdahl, North Dakota Forest Service, used with permission).
USDA Forest Service RMRS-GTR-335. 2016. 191

The resin dries into white or light blue patches (fig. 53-2A). There is little external evidence of the canker margin, but it can be found by exposing the inner bark. Infected bark 

tissue and cambium are brown in contrast to the normal cream color of healthy tissue. 

The wood beneath infected bark is not discolored (fig. 53-2B). Branches that died from 

other causes may be colonized and contain fruiting bodies of V. kunzei, but resin rarely 

exudes from such branches. Resin often exudes from wounds.

Asexual fruiting bodies (pycnidia) of the fungus develop in infected bark, but are 

usually not visible on the bark surface. Gentle scraping of cankered bark will expose the 

small (1 to 3 mm diameter) black pycnidial stromata (fig. 53-2B). Yellow or orange spore 

masses or tendrils may exude from the pycnidia during wet weather. The spores (conidia) 

are clear (hyaline), sausage shaped (allantoid), one-celled, and 4 to 6 μm × 1 μm. Sexual 

fruiting bodies (perithecia) are sometimes associated with pycnidia, but they are usually 

on branches that have been dead for several years. Perithecia are smaller than pycnidia 

and are grouped in black stromata. Ascospores from perithecia are hyaline, allantoid, onecelled, and 5 to 9 μm × 1.5 μm.

Disease Cycle

Conidia are released during all seasons except winter, whereas ascospores are released 

only in spring and early summer. Both the conidial and ascospore stages are infective. 

Spores from fruiting bodies on cankered branches are spread to branches on the same or 

other trees by rainsplash, wind, humans, and likely insects and birds. Experimental infection has succeeded only through wounds. The fungus kills bark as it grows, and usually 

girdles infected branches and stems one or more years after infection. Cankers on girdled 

branches expand toward the stem until the entire branch is dead. Fruiting bodies form in 

infected bark. The fungus overwinters as fruiting bodies and mycelium in cankered bark, 

and spores are exuded from those fruiting bodies to complete the disease cycle.

Figure 53-2—Valsa canker on blue spruce. (A) Resin from cankered branches, readily 

visible on dead bark a few months after infected needles are cast. (B) Black fruiting bodies 

(pycnidia) of V. kunzei (white arrows) in brown cankered inner bark of infected spruce 

branches, as revealed by gentle scraping. Wood beneath cankers is not discolored (black 

arrow) (A, B: James A.Walla, Northern Tree Specialties, used with permission).
192 USDA Forest Service RMRS-GTR-335. 2016.

Damage

Blue spruce is the most severely damaged host in the Great Plains, and serious damage 

is more common in the northern United States and in Canada. In most areas, branch cankering is the predominant damage. Cankers may develop on or grow into stems and result 

in top dieback, particularly on Norway spruce in some other areas. Damage may occur 

in ornamental, plantation, and windbreak settings. The disease destroys the symmetry of 

spruce trees; reduces their effectiveness in blocking wind, snow, and noise; and in time, 

reduces their vigor. Valsa canker rarely kills blue spruce but has the same effect when 

the trees are removed because of the damage. Damage is increasingly serious as trees get 

older and larger but may occur on trees less than 10 years old if they are stressed or near 

infected larger trees.

In some areas, Valsa canker is thought to damage only trees weakened by environmental stress, especially drought. In the northern Great Plains, Valsa canker commonly infects 

apparently healthy trees, but is more common and spreads more quickly on less vigorous 

trees.

Management

Healthy trees are less susceptible to damage, so trees should be managed for good 

vigor. If possible, water and fertilize trees early in the growing season as needed. Reduce 

chances of infection by preventing wounds and by maintaining good air circulation 

around the trees. Do not bring potentially infected branches into the area (for example, 

by discarding or storing spruce boughs or logs near healthy trees). Blue spruce are most 

susceptible to damage in the Great Plains, so consider planting other species or varieties 

if Valsa canker is a threat.

If trees become infected, prune out all diseased and dead branches and destroy or 

dispose of them offsite. Either prune to the trunk, or if only a small portion of a branch 

is affected, determine the proximal border of the canker and prune to the nearest healthy 

lateral branch beyond the canker. Major pruning is best done in late winter before spores 

are released, but may be done during dry periods whenever dead branches are found. 

It is critical to be aware that there will be asymptomatic cankers during the first round 

of pruning, so at least two rounds of pruning are required before prior infections are 

removed (suggested times are late winter and about one month after budbreak). Two 

consecutive years of thorough sanitation will substantially reduce future infection, but 

trees should be monitored annually thereafter to find and remove any additional cankered 

branches. Disinfect pruning tools with rubbing alcohol or 10 percent bleach after each 

cut. Protective and systemic fungicides have been recommended or labeled for control of 

Valsa canker on spruce, but efficacy information is not generally available.

Selected References

Proffer, T.J.; Hart, J.H. 1994. Wound inoculations of Colorado blue spruce with 

isolates of Leucostoma (Cytospora) kunzei and other Cytopsora species. Journal of 

Arboriculture. 20: 215–221.

Sinclair, W.A.; Lyon, H.H. 2005. Diseases of trees and shrubs. 2nd ed. Ithaca, NY: 

Cornell University Press. 660 p.
USDA Forest Service RMRS-GTR-335. 2016. 193

54. Western Gall Rust of Pines

James T. Blodgett and Kelly S. Burns; revised from Glenn W. 

Peterson and James A. Walla (Riffle and Peterson 1986)

Peridermium (Endocronartium) harknessii is a rust pathogen that causes woody swellings (galls) and cankers that deform branches and stems of hard pines (Pinus spp.). A less 

common name for this disease is pine-to-pine rust because unlike many rust diseases, it 

has no alternate host and therefore spreads directly from pine to pine.

Hosts and Distribution

The fungus infects many native hard pines including ponderosa (P. ponderosa), 

lodgepole (P. contorta), and jack (P. banksiana) pines. It also infects the exotic Scots 

(P. sylvestris) and mugo (P. mugo) pines. Western gall rust is found throughout pine 

forests of western and northern North America, in the northern Great Lakes region, and 

in scattered locations in eastern North America. It is more common in the western Great 

Plains, but can occur throughout the region. The western gall rust fungus often causes 

damage in nurseries, plantations, and landscape plantings, and can cause damage in 

natural stands.

Symptoms and Signs

Western gall rust is characterized by round or pear-shaped galls on branches 

(fig. 54-1), and rounded or targetlike cankers on stems (fig. 54-2). Galls are most 

conspicuous in spring and early summer when the surface ruptures, exposing bright 

yellow-orange spores (fig. 54-3). Witches’-brooms (bushy masses of branches) sometimes develop in affected trees (fig. 54-4).

Figure 54-1—Branch galls of western gall rust: (A) Round, and (B) Pear shaped (Joseph O’Brien, 

U.S. Forest Service, Bugwood.org (left) and U.S. Forest Service, Bugwood.org).

Figure 54-2—Cankers caused by western gall rust: (A) Swollen stem cankers in ponderosa pine, 

and (B) Sunken targetlike canker in lodgepole pine (James T. Blodgett, U.S. Forest Service).

A

A

B

B
194 USDA Forest Service RMRS-GTR-335. 2016.

Disease Cycle

White to orange pustules (aecia-like1) full of yellow-orange spores (aeciospore-like1) 

form on galls or, less commonly, at the edges of stem cankers in late spring or early 

summer. The aecia-like pustules rupture, releasing spores that disperse by wind in May 

and June. Spores germinate during moist, cool weather and infect expanding needles 

and young elongating shoots of pines. Infection rarely occurs through wounds on older 

branches and stems. Stem cankers are usually initiated by infected side branches or 

branch galls adjacent to the main stem (fig. 54-5). Mass infections tend to occur in wave 

years when weather is favorable for infection. Galls form mostly in the summer following 

the year of infection. Sporulation typically occurs in the second or third year after infection, and can continue for a few years. The rust pathogen stimulates rapid division of host 

cambial cells, which results in gall formation.

Damage

Western gall rust affects pines of all ages, causing growth loss, branch death, deformity, and occasionally mortality. Mortality is most common in seedlings and saplings 

as galls can quickly girdle small stems. Abundant galls on branches may cause tree 

stunting. Pines in landscape and Christmas tree plantings may lose value owing to 

1 Aecia-like and aeciospore-like indicate morphology that parallels typical demicyclic heteroecious 

rusts. The terms aecia and aeciospore do not describe the life stages of typical Peridermium species.

Figure 54-3—Branch gall of western gall rust sporulating on lodgepole pine (James T. Blodgett, U.S. Forest Service).

Figure 54-4—Witches’-brooms in Scots pine branches caused 

by western gall rust (Robert L. Anderson, U.S. Forest Service, 

Bugwood.org).
USDA Forest Service RMRS-GTR-335. 2016. 195

Figure 54-5—Stem canker that was initiated by a branch 

gall of western gall rust (James T. Blodgett, U.S. Forest 

Service).

stunting, witches’-brooms, and 

branch death. Stem cankers can 

severely deform larger trees, 

resulting in wind breakage or 

reduced merchantable volume, 

but they seldom girdle large trees. 

Uninfected trees of the same 

species are often common near 

similar-aged trees with many 

galls or cankers, or both. This is 

likely due to genetic variation in 

resistance to the disease in the 

population.

Management

Management of western gall 

rust can be difficult given the lag 

between infection and symptom 

development. Management 

options that can reduce disease 

impacts include sanitation, removal of the infection source, using disease-free planting stock, planting resistant hosts 

or nonhost species, and fungicide treatments.

Sanitation by removal of all trees with infections and careful inspection of remaining 

trees is often the only option in established stands. Eradication of the fungus by gall 

removal is not feasible in mature trees. In areas where western gall rust incidence is high, 

it may be necessary to replant with resistant species. Trees with stem cankers can be hazardous in recreation areas and should be given priority for removal.

In nurseries, the most effective and economical management method is the removal 

of nearby sources of infection. Gall-bearing pines within 0.5 mile of nurseries should be 

pruned free of all galls or felled. Destroy gall-bearing seedlings before sporulation.

Prevention is the best method of management in field plantings. Examine trees for 

galls before planting and again the following two springs. Obtain planting stock only 

from a nursery that is free of western gall rust.

The use of resistant hosts or nonhost species can substantially reduce or eliminate 

damage from western gall rust. Ponderosa pine from different geographic sources display 

considerable variation in susceptibility to this rust pathogen. Some highly resistant 

sources have been identified, but commercially certified, resistant stock may be difficult 

to find. When planting in areas where western gall rust is known to be a problem, use 

nonhost species that are adapted to the site.

In some states, fungicide treatments can be used for Christmas trees and high-value 

landscape trees. Consult your state or local extension agent regarding appropriate registered fungicides.

Selected References

Hiratsuka, Y.; Powell, J.M. 1976. Pine stem rusts of Canada: identification, hosts, 

distribution, morphology, life cycles, cytology, damage, epidemiology, and control. 

Tech. Rep. 4. Edmonton, AB: Canadian Forestry Service. 83 p.

Moltzan, B.D.; Blenis, P.V.; Hiratsuka, Y. 2001. Effects of spore availability, spore 

germinability, and shoot susceptibility on gall rust infection of pine. Plant Disease. 85: 

1193–1199.

van der Kamp, B. 1994. Lodgepole pine stem diseases and management of stand density 

in the British Columbia interior. The Forestry Chronicle. 70: 773–779.

Walla, J.A.; Stack, R.W. 1979. Western gall rust in North Dakota. Plant Disease Reporter. 

63: 432–433.
196 USDA Forest Service RMRS-GTR-335. 2016.

55. Pine Wilt

Timothy C. Todd and Mark O. Harrell

Pine wilt is caused by the pinewood nematode (PWN), Bursaphelenchus xylophilus, 

a microscopic roundworm (fig. 55-1) that feeds on the epithelial cells that line the resin 

canals. The nematode also feeds on fungi that proliferate in the wilting tree and in the 

pupal chambers of its beetle vector, pine sawyers of the genus Monochamus (fig. 55-2). 

Originally described from specimens collected from longleaf (Pinus palustris) and 

shortleaf (P. echinata) pines in Texas, Louisiana, and Virginia in 1934, PWN was first 

identified (as B. lignicolus) as the causal agent of pine wilt in Japanese pine forests in 

1971. In 1979, PWN was reported as the causal agent of pine wilt disease in Austrian 

(P. nigra) and Scots (P. sylvestris) pines in Missouri. The disease has caused extensive 

losses to these exotic pine species in the Midwest and Great Plains.

Hosts and Distribution

B. xylophilus has been isolated from 

numerous species of pine, and occasionally from other conifer species, including 

larch (Larix spp.), fir (Abies spp.), and 

spruce (Picea spp.). Pine wilt, with 

PWN as the primary agent, has not been 

documented for conifers other than pines, 

however, and many of these, including 

Norway (P. abies) and blue (P. pungens) 

spruce, balsam fir (A. balsamea), and 

Douglas-fir (Pseudotsuga menziesii), are 

recommended as replacement trees for 

Scots and Austrian pines in the Midwest. 

Some uncertainty also remains about 

the host status and susceptibility of 

individual pine species, but species in 

the Australes subsection (for example, 

loblolly [P. taeda], longleaf, pitch 

[P. rigida], and slash [P. elliottii]) generally are highly resistant. In contrast, 

species in the Pinus subsection (for 

example, Austrian, Japanese black 

[P. thunbergii], mountain [P. uncinata], 

and Scots) generally are highly susceptible. In the Midwest, pine wilt most 

commonly is associated with Scots and 

Austrian pines, with the primary impact 

on Scots pine in landscape plantings and 

windbreaks (fig. 55-3). Categorization 

of resistance and susceptibility in pine 

species is further confounded by variability in host preference, pathogenicity, 

and molecular phenotype among PWN 

populations in both the United States and 

Japan. Virulent and avirulent pathotypes 

appear to be widespread and there is evidence for interbreeding between them.

Figure 55-1—Pinewood nematodes: (A) Female, 

and (B) Male adult. Insert shows the distinctive 

shape of the male spicule, which is characteristic 

of the species (Timothy C. Todd, Kansas State 

University, used with permission).

A B

Figure 55-2—Pine sawyer (A) larva, and (B) 

adult beetle ((A: Mark O. Harrell, Nebraska Forest 

Service, University of Nebraska, used with permission; B: Laurie Stepanek, Nebraska Forest Service, 

used with permission).

A

B
USDA Forest Service RMRS-GTR-335. 2016. 197

The PWN is widely distributed throughout the United States, but incidence of pine 

wilt is concentrated in Iowa, Illinois, Missouri, Kansas, Oklahoma, and Nebraska. 

Isolated outbreaks of pine wilt associated with the transport of infested firewood have 

occurred in western Kansas, and the disease recently has been reported on Scots pine 

in Colorado. Presence of PWN is not always indicative of pine wilt, however, as the 

nematode can be introduced into dead or dying trees during vector oviposition, and as 

discussed above, avirulent pathotypes are known to occur.

Studies of the genetic diversity among PWN isolates have concluded that the 

nematode originated in North America. It was introduced into Japan early in the 20th

century, and subsequently spread to China, South Korea, and, more recently, Portugal. 

B. xylophilus and its vectors are listed as A1 quarantine pests by the European and 

Mediterranean Plant Protection Organization (EPPO). 

Symptoms and Signs

Reduction in oleoresin flow and rapid tree death are the characteristic symptoms of 

pine wilt. Initially needles on affected branches change in color to faded green, then to 

light brown (fig. 55-4). The discoloration may be restricted to individual branches at first, 

but it progresses rapidly, often spreading to the entire tree within a few weeks. Trees often 

die between midsummer and fall, but death can occur at other times of the year. Trees 

beginning to show symptoms in fall may not become completely brown until after winter, 

when warmer weather returns. 

Rapid death of pines can also be caused by environmental stresses. The presence of 

PWN can be confirmed by clinical diagnosis of wood samples collected from the base of 

lower limbs soon after symptoms appear.

Disease Cycle

The life cycles of the PWN and its beetle vectors are closely synchronized. The most 

widespread and efficient vectors of PWN in the United States are the nearly identical 

Carolina (M. carolinensis) and Southern (M. titillator) pine sawyer and the white-spotted 

sawyer (M. scutellatus). The nematode may be transmitted to healthy trees during maturation feeding by newly emerged beetles or to dead or stressed trees during oviposition 

by mature beetles. Transmission efficiency during M. carolinensis feeding has been 

demonstrated to approach 50 percent of fed-upon branches for mature Scots pines under 

field conditions. Actual numbers of transmitted nematodes are correlated with numbers of 

nematodes carried by the beetle but generally appear to be low; infection results from the 

introduction of a small number of nematodes through one or more feeding wounds.

Figure 55-3—Pine wilt in Scots pine windbreak (Mark O. Harrell, Nebraska Forest Service, University 

of Nebraska, used with permission).
198 USDA Forest Service RMRS-GTR-335. 2016.

 The progression of wilt symptoms is accompanied by the emission of volatiles that 

attract mature beetles to diseased trees, where mating and oviposition occur. Proliferation 

of fungi in the dying tree provides not only a secondary food source for PWN populations 

that previously fed upon living plant tissues but also a primary food source for any nematodes that may have been transmitted during oviposition. Pupal chambers, in particular, 

harbor a diverse assemblage of microorganisms ranging from blue-stain fungi, which are 

disseminated by Monochamus and other beetles and serve as an important food source 

for the nematode, to nematophagous fungi that capture and kill nematodes and have the 

potential to reduce the number of nematodes carried by beetles emerging from the wood.

Desiccation and nematode overpopulation in the wood eventually induce the formation 

of the third juvenile dispersal stage of the nematode, which aggregates around the pupal 

chambers of vector beetles. Emergence of beetles from the pupal chamber is associated 

with the release of volatiles that stimulate the nematodes to molt to the fourth juvenile 

dispersal stage, which then enters the spiracles and trachea of the insect. Adult beetles 

emerge throughout the spring and early summer. The life cycle of M. carolinensis is temperature dependent, resulting in two generations per year in the southern United States 

but only one generation every two years in the northern United States.

Damage

Unlike other nematode diseases of conifers, the primary impact of pine wilt is not 

on seedlings but on mature trees in landscape plantings and windbreaks. Susceptible 

Scots pines can be killed within a few weeks or months. Pathogenicity of the nematode 

is determined by rates of transmission, dispersal, and reproduction within healthy trees, 

and by the interaction of secreted and surface coat proteins with host tissue cells. Water 

deficiency is the ultimate factor in rapid tree mortality, but the proximate cause appears to 

be the induction of a hypersensitivelike response to nematode feeding in susceptible trees. 

Management

Traditional control practices have focused on sanitation involving the timely removal 

and destruction of trees killed by pine wilt. This practice eliminates the breeding habitat 

of the beetle vector and prevents nematode transmission to healthy trees. More recently, 

demonstration of effective PWN control using preventive injections has resulted in the 

emergence of several new products and injection systems. The active ingredient in these 

Figure 55-4—Symptom progression in Scots pine from (A: Mark O. Harrell, Nebraska Forest Service, 

University of Nebraska, used with permission; B: Nebraska Forest Service, University of Nebraska, 

used with permission).

A B
USDA Forest Service RMRS-GTR-335. 2016. 199

products is abamectin, a natural fermentation product of the soil bacterium Streptomyces 

avermitilis with nematicidal properties (enhanced gamma-aminobutyric acid [GABA] 

activity). Abamectin injections in PWN-inoculated, 13- to 20-foot-tall Scots pine trees 

have been observed to increase the survival rate of treated trees to 75 percent, compared 

to 25 percent in untreated control trees, up to one year after injection. Similarly, abamectin injections in mature trees exposed to a natural epidemic increased the survival rate 

of treated trees to 96 percent, compared to 33 percent in untreated control trees, up to 

two years after injection. Curative injections after symptoms have started appearing, in 

contrast, are ineffective. Regardless of the efficacy of preventive injections, avoid new 

plantings of Scots and Austrian pines wherever possible.

Wood chips from PWN-infested trees can be used as mulch, but should be used only 

around nonsusceptible tree species and should not be placed against fresh wounds on 

pines. Research has shown that the nematode can survive in infested chips for up to 

12 weeks and is capable of infecting wounded roots and stems that are in contact 

with the mulch.

Selected References

Akbulut, S.; Stamps, W.T. 2012. Insect vectors of the pinewood nematode: a review of 

the biology and ecology of Monochamus species. Forest Pathology. 42: 89–99.

Bolla, R.; Winter, R.E.K.; Fitzsimmons, K.; Linit, M.J. 1986. Pathotypes of pinewood 

nematode Bursaphelenchus xylophilus. Journal of Nematology. 18: 230–238.

Futai, K. 2013. Pine wood nematode, Bursaphelenchus xylophilus. Annual Review of 

Phytopathology. 51: 61–83.

Halik, S.; Bergdahl, D.R. 1992. Survival and infectivity of Bursaphelenchus xylophilus in 

wood chip-soil mixtures. Journal of Nematology. 24: 495–503.

James, R.F.; Tisserat, N.A.; Todd, T.C. 2006. Prevention of pine wilt of Scots pine (Pinus 

sylvestris) with systematic abamectin injections. Arboriculture & Urban Forestry. 32: 

195–201.

Linit, M.J. 1990. Transmission of pinewood nematode through feeding wounds of 

Monochamus carolinensis (Coleoptera: Cerambycidae). Journal of Nematology. 22: 

231–236.

Wingfield, M.J. 1987. Fungi associated with the pine wood nematode, Bursaphelenchus 

xylophilus, and cerambycid beetles in Wisconsin. Mycologia. 79: 325–328.
200 USDA Forest Service RMRS-GTR-335. 2016.

56. Root Parasitic Nematodes in Junipers and Pines

Timothy C. Todd and Jon A. Appel; revised from Glenn W. Peterson 

and Jerry W. Riffle (Riffle and Peterson 1986)

The root-lesion nematode Pratylenchus penetrans was the most frequently encountered plant-parasitic nematode in a 1960 survey of Nebraska tree nurseries. Damage from 

this nematode was reported for Austrian (Pinus nigra) and ponderosa (P. ponderosa) 

pine seedlings, but was not observed for windbreak stands of pines and eastern redcedar 

(Juniperus virginiana). In a 2004 to 2006 survey of windbreaks in Kansas1, root-lesion 

nematodes were recovered from 25 percent of pine stands and 35 percent of eastern redcedar stands (table 56-1), but, again, damaging population densities were not observed.

Hosts and Distribution

Pratylenchus penetrans (fig. 56-1) is a cosmopolitan pest in North America, with a 

host range that encompasses grain crops, vegetables, fruit trees, and woody ornamentals. 

In central Nebraska, this nematode was recovered from roots of eastern redcedar, Rocky 

Mountain juniper (J. scopulorum), white (Picea glauca) and blue (P. pungens) spruce, 

Austrian pine, and ponderosa pine.

Nematodes commonly associated with conifers in Kansas windbreaks include 

pin (Paratylenchus) and spiral (Helicotylenchus) for eastern redcedar, and dagger 

(Xiphinema) and spiral for Austrian and ponderosa pines (table 56-1). These nematodes 

also were prevalent in hardwood stands (see chapter 40).

Table 56-1—Prevalence and abundance of nematode genera recovered from 

eastern redcedar and pine stands during a three-year survey of 81 Kansas 

windbreaks

Mean (maximum) number

Nematode Prevalence (percent) of nematodes/100 cm3 soil

common name Redcedar Pine Redcedar Pine

(genus) (n=46) (n=32) (n=46) (n=32)

Dagger 24 50 7 (60) 16 (130)

(Xiphinema)

Lance 0 0 0 (0) 0 (0)

(Hoplolaimus)

Pin 57 38 103 (1,510) 34 (590)

(Paratylenchus)

Ring 2 16 0 (10) 3 (50)

(Mesocriconema)

Root-lesion 35 25 12 (90) 6 (50)

(Pratylenchus)

Spiral 46 53 31 (420) 45 (520)

(Helicotylenchus)

Stunt 28 34 7 (100) 8 (90)

(Tylenchorhynchus)

Stubby-root 0 3 0 (0) 0 (10)

(Paratrichodorus)

1Data on file with the Kansas Department of Agriculture, Topeka, KS.
USDA Forest Service RMRS-GTR-335. 2016. 201

Symptoms and Signs

Aboveground symptoms of nematode damage 

are most noticeable on nursery seedlings and typically manifest as irregular patches of stunted plants, 

often accompanied by signs of nutrient deficiency. 

Symptoms on roots are mostly nonspecific and may 

consist of formation of woody swellings (galling), 

restricted root development, and root necrosis. Rootlesion nematodes may produce necrotic lesions in 

the cortical tissue of feeder roots and can predispose 

plants to root rots and other disease complexes. 

Nematode extraction from soil and roots is always 

necessary to confirm a diagnosis of nematode 

damage.

Disease Cycle

Root-lesion nematodes are migratory endoparasites that enter and feed in the cortex of 

the feeder roots of host trees. These nematodes hatch as second-stage juveniles from eggs 

laid in or near host roots and develop through two additional juvenile stages into the adult 

stage, typically within four to six weeks, depending on temperature and moisture conditions. Migration between roots and soil can occur continuously, but movement into the 

soil environment is most pronounced during host senescence.

Dagger nematodes are ectoparasites with elongate stylets that facilitate prolonged 

feeding on root cortical and stelar tissues. The life cycle of dagger nematodes is 

longer than that of root-lesion nematodes, requiring one to two years for completion. 

Reproduction and population increase also are slower; nonetheless, perennial hosts such 

as trees provide ample time for damaging populations to develop.

Damage

Root-parasitic nematode damage in conifers has been observed primarily for seedlings 

in nurseries. Root-lesion nematodes were associated with extensive damage to redcedar seedlings in a Nebraska nursery, but were not associated with obvious damage to established 

pines, junipers, or spruce in windbreaks in Kansas and Nebraska. Nematodes known 

to damage conifer seedlings outside of the Great Plains include the needle nematode 

Longidorus americanus and the stunt nematode Tylenchorhynchus ewingi in pine 

nurseries in the southern United States, and P. penetrans and the dagger nematode 

X. bakeri in spruce and fir nurseries in the Pacific Northwest.

X. americanum population densities above 100 individuals/100 cm3 (6 cubic inches) soil were associated with poor health of pines in Kansas windbreaks. Similarly high 

population densities of this nematode also were indicative of poor tree health in hardwood windbreak stands (see chapter 40).

Management

Nematode management options for windbreaks are limited. Weed control may restrict 

nematode population size, and cultural practices that promote tree health may increase 

tolerance to nematode feeding pressure. Combine routine fumigation of seedling production areas with sanitation practices and rotation with nonhost cover crops to avoid 

damage in nurseries. Most soil fumigant alternatives to methyl bromide have been shown 

to provide adequate control of plant-parasitic nematodes, but all fumigants provide only 

temporary control of nematodes and must be used routinely.

Figure 56-1—Adult female 

Pratylenchus penetrans (Timothy C. 

Todd, Kansas State University, used 

with permission).
202 USDA Forest Service RMRS-GTR-335. 2016.

Selected References

Cram, M.M.; Fraedrich, S.W. 2012. Nematode damage and management in North 

American forest nurseries. Tree Planters’ Notes. 55: 27–35.

McElroy, F.D. 1990. Nematodes: Pratylenchus penetrans; Xiphinema bakeri. In: Hamm, 

P.B.; Campbell, S.J.; Hansen, E.M., eds. Growing healthy seedlings: identification and 

management of pests in Northwest forest nurseries. Special Publ. 19, Corvallis, OR: 

Oregon State University: 62–65.

Peterson, G.W. 1962. Root lesion nematode infestation and control in a plains forest tree 

nursery. Res. Note RM-75. Fort Collins, CO: U.S. Department of Agriculture, Forest 

Service, Rocky Mountain Forest and Range Experiment Station. 2 p.

Peterson, G.W. 1970. Response of ponderosa pine seedlings to soil fumigants. Plant 

Disease Reporter. 54: 572–575.

Ruehle, J.L. 1973. Nematodes and forest trees—types of damage to tree roots. Annual 

Review of Phytopathology. 11: 99–118.
USDA Forest Service RMRS-GTR-335. 2016. 203

Glossary (with plurals in parentheses)

ABAXIAL: referring to the bottom surface of a leaf.

ACERVULUS (ACERVULI): a small subcuticular or subepidermal cushionlike asexual 

fruiting body, without a covering of fungus tissue, producing conidia in a mass, which 

escape through a break in the host tissue.

ACIDIC: having a pH below 7: containing or having the properties of an acid.

ADAXIAL: referring to the upper side of the leaf. 

AECIOSPORE: one of several kinds of spores produced by a rust fungus. Formed in and 

released from a fruiting structure called an aecium.

AECIUM (AECIA): a cuplike fruiting structure produced by certain rusts, in which 

chains of spores (aeciospores) are developed.

AGAR: a substance from certain red algae used to make culture media into gels upon 

which organisms are grown.

ALKALINE: characterized by high pH. Soils with pH levels above 7.0 are considered 

alkaline; greater than 8.5 are considered highly alkaline.

ALLANTOID: slightly curved with rounded ends; sausagelike in form.

ALPHA-SPORE: a fertile spore of the asexual stage of the Diaporthaceae (Phomopsis) 

that is fusiform to oblong and biguttulate (having two oil droplets inside the cell); a 

beta-spore, usually hook shaped, may be produced in addition.

ALTERNATE HOST: one or the other of the two unlike host plants parasitized by a 

heteroecious (involving two hosts) fungus such as a typical rust fungus, for example, 

either the juniper or apple host of the cedar-apple rust fungal disease complex.

ANAEROBE: a microorganism that can live and grow where there is no free oxygen.

ANAMORPH: the asexual form of a fungus (the form that is characterized by 

conidiomata).

ANNULUS: a ringlike partial veil around the stem of a fruiting body after expansion of 

the pileus (cap).

ANTHRACNOSE: a type of plant disease that typically is a leaf and twig blight. 

Common on many hardwoods.

APOTHECIUM (APOTHECIA): a cuplike or saucerlike sexual fruiting body that 

produces ascospores.

ARCHAEA: a kingdom of single-celled prokaryotic microorganisms, once considered 

bacteria, that inhabit a broad range of habitats.

ASCOMYCETE: fungi belonging to the Ascomycota, the largest group of fungi and for 

which the ascus is the main diagnostic characteristic. 

ASCOSPORE: a spore produced in an ascus (see ASCUS).

ASCUS (ASCI): a saclike cell of the perfect (sexual) stage of the Ascomycetes in which 

ascospores (usually eight) are produced.

ASEXUAL STAGE: a stage in the life cycle of a fungus in which spores are produced 

without a previous sexual fusion. Also called IMPERFECT STAGE.

BASIDIOMYCETE: any fungus in the phylum Basidiomycota producing spores on a 

basidium.

BASIDIOCARP: spore-producing structure of basidiomycetous fungi. 

BASIDIOSPORE: the spore produced by the sexual stage of the Basidiomycetes.

BASIDIUM (BASIDIA): a cell, usually terminal, in which nuclear fusion and meiosis 

occur and each of the four haploid nuclei pass into one of four forming spores.

BETA-SPORE: see ALPHA-SPORE.

BLIGHT: a general term for a plant disease causing rapid death or dieback.

BRASH: the characteristic that causes a piece of wood to break abruptly across the grain, 

with relatively low resistance to such breaking. 
204 USDA Forest Service RMRS-GTR-335. 2016.

BROOM: see WITCHES’-BROOM.

BROWN ROT: a type of wood decay caused by fungi that degrade cellulose but not 

lignin, do not produce extracellular phenoloxidases, and generally give negative 

oxidase tests. Decayed wood appears brown, fractures into cubical pieces, and is 

crumbly (not to be confused with the disease Brown Rot of Stone Fruits).

CALLUS: undifferentiated cell formation at the site of injured tissue. Note: the term is often 

used incorrectly to refer to wound closure with woody tissue. See WOUNDWOOD.

CANKER: a definite, relatively localized necrotic lesion primarily of the bark and 

cambium.

CELLULOSE: long chains of sugar molecules and the main component of the plant cell 

wall giving wood its mechanical strength.

CHASMOTHECIUM: see CLEISTOTHECIUM.

CHLOROSIS: an abnormal yellowing of the foliage.

CHLOROTIC: abnormally yellow.

CIRRHUS (CIRRUS, CIRRHI): tubelike mass of spores exuded from a fungal fruiting 

structure. Also called a TENDRIL or sporehorn.

CLAMP CONNECTION: a hyphal outgrowth formed by certain fungal species 

functioning to allow the transfer of fungal nuclei.

CLEISTOTHECIUM (CLEISTOTHECIA): a closed fruiting body, without an opening, 

containing asci.

CONIDIOGENOUS CELL: a cell from or within which a conidium is directly produced.

CONIDIOMA (CONIDIOMATA): A specialized multihyphal, conidia-bearing structure.

CONIDIOPHORE: a specialized hypha-bearing conidium.

CONIDIUM (CONIDIA): a spore formed asexually, usually at the tip or side of a hypha.

CONK: a type of fruiting (spore-forming) structure, usually formed by certain wood 

decaying fungi in the Basidiomycetes. It is often bracketlike and is also referred to 

as “sporocarp,” “sporophore,” “FRUITING BODY,” “carpophore,” “fructification,” 

“basidiocarp,” “punk,” “bracket,” and “shelf.”

CONTEXT (of CONK): the inner tissue of the pileus (cap), that is, the tissue lying 

between the upper surface and the tube or pore layer. Sometimes designated as the 

“trama” of the pileus.

COVER CROP: a crop, natural or introduced, that is grown alternately with the main 

crop. Used to prevent erosion, improve soil characteristics, and to favor depletion of 

plant pathogenic fungus and nematode populations.

CULTIVAR: a group of cultivated plants distinguished by any useful, reproducible 

character; a cultivated variety.

CUTTING: detached portion of stem or other plant part that, when rooted, produces a 

whole plant.

DAEDALEOID-PORES: pores with elongated and sinuous mouths.

DECAY: the decomposition of plant tissue by fungi and other microorganisms.

DEFOLIATION: loss of current year’s or past years’ foliage.

DELIQUESCENT: lack of terminal dominance on branch ends resulting in terminal and 

lateral twigs that grow about the same amount annually, often restricted to one or a 

few branches or to sprouts along the trunk.

DEMICYCLIC: Rust life cycle lacking the uredial (repeating) stage.

DEXTRINOID: stained yellowish- or reddish-brown by Melzer’s iodine.

DESSICATE: to dry (dehydrate) thoroughly. 

DIEBACK: the death of parts of a tree or plant, usually from the top downward.

DIMITIC: referring to fungi that have two types of hyphae.
USDA Forest Service RMRS-GTR-335. 2016. 205

DISEASE: unfavorable change of the function or form of a plant from normal; caused by 

a pathogenic agent or unfavorable environment.

DISEASE CYCLE: the chain of events in the development of a disease.

DISSEPIMENTS: the pore walls of polypores.

DUPLEX: (of a context of a sporocarp) in two layers. 

ECTOPARASITE: a parasite living outside of its host.

EFFUSED: stretched out flat.

EFFUSED-REFLEXED: spread out over a substratum and turned back at the margin to 

form a pileus (cap).

ELLIPSOID: In the shape of an ellipse.

ENDOPARASITE: a parasite living inside of its host.

ENDOPHYTIC: describes an organism (endophyte), usually a fungus or bacteria, that is 

able to live within a plant for at least part of its life cycle without causing disease.

EPIDEMIC: pertaining to a disease that has built up rapidly in a population or area and 

reached injurious levels.

EPIDERMIS: the outermost layer of cells on the primary plant body.

EPIPHYTE: an organism that lives nonparasitically on the surface of a plant.

ERADICANT: a fungicide, applied to foliage of a plant, that has a direct effect upon 

organisms which have already invaded the host.

EXUDATE: matter that oozes out or is secreted.

FACULTATIVE PARASITE: an organism that is normally saprophytic but that is capable 

of living as a parasite only when unfavorable conditions predispose the host so that it 

is unusually susceptible.

FALLOW: referring to cultivated land allowed to lie idle or unplanted during the growing 

season.

FASCICLE SHEATH: a sheath around the base of a cluster or bundle of needles.

FEEDER ROOTS: succulent, actively growing rootlets of plants.

FLAGELLUM (FLAGELLA): a whiplike part of some cells, especially of certain 

bacteria, that is an organ of locomotion.

FOLIAGE: leaves of a plant or tree.

FRUITING BODY: any of a number of kinds of reproductive structures that produces 

spores.

FUNGICIDE: chemical that is toxic to fungi.

FUNGUS (FUNGI): any of a number of organisms considered by some authorities to be 

lower plants which lack chlorophyll.

FUSIFORM: spindlelike, narrowing toward the end.

GALL (BURL): a pronounced swelling on a woody plant caused by certain insects and 

disease organisms.

GIRDLE: to destroy or remove the tissue, particularly living tissue in a rough ring around 

a stem, branch, or root.

GLABROUS: smooth, not hairy.

GLOBOSE: spherical or almost so.

GUMMOSIS: the exudation of gummy substances.

HAUSTORIUM (HAUSTORIA): absorbing organ originating on a hypha of a parasite 

and penetrating into a cell of the host.

HEARTWOOD: region at the center of woody plants mainly composed of dead cells, 

typically darker and harder than the SAPWOOD. 

HEMICELLULOSE: polysaccharide component of plant cell walls.
206 USDA Forest Service RMRS-GTR-335. 2016.

HETEROECIOUS: Describing rust fungi requiring two unrelated hosts to complete their 

life cycle.

HIRSUTE: having long hairs.

HISPID: having hairs or bristles.

HOST: The organism on or in which a pathogen exists.

HOST RANGE: all hosts that a particular pathogen attacks.

HOST-SPECIFIC: a term used to describe those pathogens that attack hosts of only a 

certain species or cultivar.

HYALINE: transparent, having no color.

HYMENIUM: the general region of a sporocarp bearing basidia, that is, the layer of 

tubes. Alternatively, the actual spore-producing layer, made up of basidia and whatever 

type of non-reproductive tissue that may be present with them and forming a layer 

lining the inside of the tubes.

HYPHA (HYPHAE): a fungal filament; collectively called mycelium.

HYSTEROTHECIUM: an elongated sexual fruiting structure splitting longitudinally to 

release spores; of ascomycetes and lichens. 

IMPERFECT STAGE: that part of the life cycle of a fungus in which only conidia and no 

sexual spores are produced. Also called ASEXUAL STAGE.

INFECT: to invade and persist in an organism potentially causing disease.

INFECTION COURT: the area in which the pathogen first established itself on or in the 

host.

INFEST: to be present within an area in such numbers as to be a disease hazard.

INOCULATE: to place an organism or substance on or in another potential host organism 

or a substratum.

INOCULUM: the organism or substance, e.g., spores of a pathogen, that can inoculate an 

organism or substratum.

INTERNODE: the portion of the stem between two nodes.

IN VITRO (or VITRO): taking place in a test tube, laboratory, or otherwise outside of a 

living organism.

IN VIVO (or VIVO): taking place in a living organism. 

JUVENILE LEAVES: foliage characteristic of the immature form of leaves for species 

that have multiple leaf forms, e.g., awl-shaped needles of junipers.

LAMELLATE: vertical plates on the underside of the pileus. 

LATENT INFECTION: an established infection that does not show its presence.

LEAF ABSCISSION: the separation of leaves from plants caused by the weakening of 

cell walls in a special layer (abscission layer) at the base of the petioles.

LEAF PETIOLE: the slender, usually cylindrical portion of a leaf, that supports the blade 

and is attached to the stem.

LEAF SPOT: a disease symptom or a type of leaf disease; typically characterized by 

distinct lesions on leaves.

LESION: a defined necrotic area.

LIFE CYCLE: in the case of diseases, the stage or series of stages between the formation 

of one form of a pathogen (e.g., spore), development of disease, and reappearance of 

the same form of the pathogen.

LIGNIN: A complex polymer that with cellulose is a major component of wood and the 

cell walls of plants, providing rigidity.

LOCULE: cavity in a stroma

MACROCONIDIA: the larger of two types of conidia by certain fungi, such as Fusarium

species.

MICROCONIDIA: the smaller of two types of conidia produced by certain fungi.
USDA Forest Service RMRS-GTR-335. 2016. 207

MICROSCLEROTIA: small, melanized (containing dark brown pigments) resting 

structures produced by some fungi.

MONOMITIC: having one type of generative hypha.

MULTISEPTATE: characterized by having several cross walls that divide hyphae or 

spores into a number of separate cells.

MUSHROOM: any of various rapid-growing, fleshy fungi that typically have a stalk 

capped with an umbrella-like top.

MYCELIUM (MYCELIA): a mass of hyphae that forms the vegetative filamentous body 

of a fungus.

MYCOPARASITIC: describing a fungus-parasitizing organism.

MYCOPLASMA-LIKE ORGANISM (MLO): an organism with apparent features of 

previously known mammal-infecting bacteria called mycoplasmas, now termed 

PHYTOPLASMAS, a group of plant-infecting bacteria distinct from mycoplasmas.

MYCORRHIZA (MYCORRHIAZE): type of fungus that engages in a symbiotic 

(sometimes weakly pathogenic) lifestyle with a vascular plant, growing on or in plant 

roots, aiding in water and nutrient uptake and receiving carbon in exchange.

NECROSIS: death of plant cells, usually resulting in darkening of the tissue.

NEMATODE: a roundworm with a long, cylindrical, unsegmented body.

NODULOSE: broad-based, blunt, wart-like. 

NONSELECTIVE HERBICIDES: herbicides used for control of all vegetation (grass and 

broadleaf plants).

OSTIOLE: small opening or pore in fungal fruiting structures where fungi release mature 

spores.

OVIPOSITION: the depositing (laying) of eggs through a specialized tubular organ 

called an ovipositor (typically in insects).

PARASITE: an organism living on or in, and nourished by, another living organism.

PATHOGEN: an organism that causes a disease.

PATHOGENIC: capable of causing a disease.

PEDUNCLE: the supporting axis of a single flower or a flower cluster.

PERENNIAL: continuing growth from year to year.

PERFECT STAGE: the stage in which the sexual spores are produced. Also called sexual 

stage.

PERIDERM: the outer protective layer in older stems, consisting of the phellogen and its 

derivative tissues, phellem and phelloderm.

PERITHECIUM (PERITHECIA): a closed flasklike sexual fruiting body formed by 

certain Ascomycetes and in which ascospores are produced.

PHLOEM: the tissue of the inner bark responsible for the transport of sugars and growth 

regulators.

PHYTOPLASMA: a cell-wall-less bacterium that lives in plant phloem tissue and certain 

insects, causing disease in both hosts, with various taxa causing many plant diseases; 

formerly known as MYCOPLASMA-LIKE ORGANISM (MLOs). 

PILEATE: having at least a portion of a conk with a sterile upper surface over a fertile 

lower surface (see PILEUS).

PILEUS (PILEI): the upper surface of a conk/sporocarp.

PLEOMORPHIC: the condition in which a fungus has two or more very different 

morphological forms.

POLYPORE: a group of fungi that form fruiting bodies with pores or tubes on the 

underside.

PORE SURFACE: the lower surface of a sporocarp in specimens mature enough to have 

a tube layer. It is the surface at which the pores open.
208 USDA Forest Service RMRS-GTR-335. 2016.

POST-EMERGENCE HERBICIDES: herbicides applied after germination and 

establishment of green plants.

PREEMERGENCE HERBICIDES: herbicides applied before seedling emergence, 

usually designed to inhibit germination.

PRIMARY INFECTION: infection of a host by primary inoculum (first inoculum of a 

season), for example, from overwintering spores.

PROTECTANT: pesticide applied to foliage of a plant in advance of a pathogen in order 

to prevent infection.

PRUNE: to remove dead or living parts from a plant to improve its form or condition.

PSEUDOTHECIUM: a sexual fruiting structure (ascocarp) containing randomly 

organized asci.

PUSTULE: a small, sometimes colored, blisterlike swelling.

PYCNIDIUM (PYCNIDIA): an asexual type of fruiting body, typically flask shaped, in 

which asexual spores or conidia are produced.

PYRIFORM: pear-shaped.

REFLEXED: describing a decay conk/sporocarp that is mostly resupinate, but with a 

small portion at the upper end that is curved outward, creating a small pileus.

RELATIVE HUMIDITY: the amount of moisture in the air as compared with the 

maximum amount that the air could contain at the same temperature, expressed as a 

percentage.

RESISTANCE: the ability of a host to retard, suppress, or prevent infection or 

colonization by a pathogen; also the functioning of attributes or processes that do so.

RESUPINATE SPOROCARP: describing a decay conk/sporocarp in which the entire 

structure lies flat on the substratum, that is, without forming a bracketlike or shelflike 

body (not having a pileus and having only a pore surface surrounded by a narrow 

margin).

RHIZOMORPH: a thick strand of vegetative hyphae in which the hyphae have lost their 

individuality; important in underground expansion of some types of root rot.

RHIZOSPHERE: the soil near a living root.

RIMOSE: cracked, or cracked by radial fissures.

ROT: see DECAY.

SAPROBE: fungus that derives nutrients from dead and decaying organisms.

SAPROPHYTE: an organism using dead organic material as food.

SAPWOOD: the wood beneath the bark that contains living tissue and functions in water 

and mineral transport and starch storage.

SCLEROTIUM (SCLEROTIA): a firm, frequently rounded, multicellular resting 

structure produced by fungi.

SECONDARY INFECTION: infection that commonly occurs when the infecting 

organism produces another crop of spores or infective bodies after a host has already 

become diseased.

SELECTIVE HERBICIDES: herbicides designed to target specific types of plants, while 

leaving the desired types of plants unharmed.

SEPTATE: having cross walls that divide hyphae or spores into a number of separate 

cells.

SEPTUM (SEPTA): a cross wall that divides a hypha or spore into two or more distinct 

cells.

SESSILE SPOROCARP: describing a decay conk that takes the form of a knob, bracket, 

or shelf. The conk has no stem or stalk, and the point of attachment to the substratum 

is typically lateral. Pileate, without a stem, and not effused; typical conk form.

SETA (SETAE): conical or lance-shaped, brown, sterile organs found in the hymenium of 

some fungal species.
USDA Forest Service RMRS-GTR-335. 2016. 209

SEXUAL STAGE: the stage in the life cycle of a fungus in which spores are produced 

after sexual fusion. Also called perfect stage.

SHEARING: cutting foliage and stems from trees with shears.

SHELTERBELT: see WINDBREAK.

SIGNS OF A PATHOGEN: any observable parts of a pathogen.

SILVICULTURE: the art of establishing, growing, and regenerating a forest.

SPERMATIUM (SPERMATIA): a nonmotile, uninucleate, sporelike male structure 

which empties its contents into a receptive female structure during a type of sexual 

reproduction known as plasmogamy.

SPERMAGONIUM (SPERMAGONIA): a structure that contains minute, sporelike 

bodies which in some cases have proved to be functional spermatia.

SPORE: a reproductive or resting structure of the fungi, other lower plants, and some 

bacteria.

SPORE HORN: a thread-like gelatinous mass of forced-out spores. Also called 

TENDRIL or CIRRHUS

SPOROCARP: structure on which spore-producing structures, such as basidia or asci, are 

borne. 

SPORODOCHIUM (SPORODOCHIA): a cushion-shaped stroma covered with 

conidiophores.

SPORULATE: to produce spores.

SPUR LEAVES: foliage characteristic of short (spur) branches.

STOMA (STOMATA): a pore in the leaf epidermis, surrounded by two guard cells, 

leading into an intercellular space within the plant.

STOMATAL PIT: the space above a stoma that is sunken below the leaf surface.

STRESS: a negative impact on tree health caused by biotic or abiotic factors leading to 

disruption of normal physiological processes supporting normal growth.

STROMA (STROMATA): a mass of fungal hyphae packed together to form a hard crust 

in or on which fruiting bodies are formed.

SULCATE ZONES: grooved regions of fungal fruiting bodies.

SUSCEPTIBLE: unable to resist attack by an organism or influence by a nonliving agent.

SYMPTOM: the evidence of disturbance in the normal development and function of a 

host plant, for example, chlorosis, necrosis, galls, brooms, and stunting.

SYNNEMA (SYNNEMATA): a specialized, spore-bearing structure composed of fused, 

elongated multihyphal conidiophores.

SYSTEMIC: able to move or be transported internally in a plant.

SYSTEMIC FUNGICIDES: chemicals and natural products that are absorbed into the 

tissues of plants and are toxic to fungi.

TELEOMORPH: the perfect (sexual) stage of a pleomorphic fungus, characterized by 

ascomata and basidiomata.

TELIOSPORE: the fourth of five potential spore stages of the rust fungi, from which 

the perfect stage of the basidium and basidiospore arises. Typically a resting, 

overwintering stage.

TELIUM (TELIA): a fruiting body or fruiting structure producing teliospores.

TENDRIL: a thread-like gelatinous mass of forced-out spores. Also called sporehorn or 

cirrhus.

TOMENTOSE: covered with soft matted hairs.

TOMENTUM: a layer of soft matted hairs on plant parts.

TRAMA: the fleshy tissue within a basidiocarp.

TRIMITIC: describing a fungus that has three types of hyphae.
210 USDA Forest Service RMRS-GTR-335. 2016.

TUBE LAYER OF SPOROCARP: a layer of vertically placed tubes attached to the lower 

surface of the context of a sporocarp. See HYMENIUM.

TYLOSES: outgrowth of cells in woody tissue that block water-conducting xylem tissue.

UNGULATE: in a form resembling a horse’s hoof.

UREDIOSPORE also UREDINIOSPORE: the third of five potential spore stages 

produced by the rust fungi in their life cycle, capable of infecting the host on which 

it was formed (so repeating stage). The stage is produced in a fruiting body called a 

uredium.

UREDIUM (UREDIA, UREDINIA): one of the types of fruiting bodies formed by the 

rust fungi in their life cycle. Urediospores are formed in this fruiting body.

VECTOR: an organism, usually an insect, that transmits a pathogen from one host to 

another.

VEGETATIVE MYCELIUM: a mass of hyphae constituting the body of the fungus and 

without spores.

VELUTINATE: covered with a thick layer of delicate hairs, velvet-like.

VERMIFORM: worm-like.

VILLOSE: covered with long thin, nonmatted hairs (villi).

VIRULENT: vigorously pathogenic.

WETWOOD: a discolored, water-soaked condition of the heartwood of some trees 

presumably caused by bacterial fermentation. Often associated with distinctive odor, 

gas production, and an exudation called slime flux.

WHITEROT: a type of wood decay caused by fungi that degrade all chief constituents of 

wood and leave a whitish or light-colored residue. Affected wood is often fibrous or 

spongy in texture.

WINDBREAK: a row or rows of trees that serve as a protection from wind.

WITCHES’-BROOM: cluster of branches, twigs, or foliage, often denser than normal, 

caused by certain pathogens (sometimes insects, or a combination of insects and 

pathogens), which is not typical of the host’s normal pattern of growth.

WOUNDWOOD: differentiated and organized tissue, high in lignin, that is produced by 

woody plants to cover wounds. Woundwood replaces callus tissue at the site of injury.

ZONATE: having zones characterized by different qualities.
USDA Forest Service RMRS-GTR-335. 2016. 211

Index to Host Plants With Scientific Equivalents

Host Article Number

Aigeiros, Populus section. 9, 23

Ailanthus, Ailanthus altissima (Mill.) Swingle. 16

Alder, Alnus spp. 25

Apple, Malus spp. 5, 8, 17, 22, 26, 36, 38, 39, 52

Apricot, Prunus spp. 4, 14, 17, 18

Arborvitae, Thuja spp. 1, 36, 42, 47, 50

Aronia (see Chokeberry)

Ash, Fraxinus spp. 1, 2, 6, 8, 11, 19, 30, 36, 37, 40

Black, F. nigra Marsh. 37

Green, F. pennsylvanica Marsh. 11, 30, 37, 40

White, F. americana L. 11, 37

Aspen, Populus spp. (see also Poplar) 5, 9, 10, 23, 24, 27, 30, 32, 38, 39

Bigtooth, P. grandidentata Michx. 27

Quaking, P. tremuloides Michx . 9, 10, 23, 27

Baldcypress, Taxodium distichum (L.) Rich. 5

Basswood, Tilia spp. (see also Linden) 2, 8, 15, 19, 39

American, T. americana L. 2, 8, 15, 19, 39

Beech, Fagus spp. 36, 39

Birch, Betula spp. 1, 5, 8, 30, 36, 38, 39

Paper, B. papyrifera Marsh. 38

Bittersweet, Celastrus spp. 19

Black Locust, Robinia pseudoacacia (see also Locust). 30, 36, 39

Buckthorn, European, Rhamnus cathartica L. 19, 30

Buffaloberry, Shepherdia spp. 19, 30

Butternut, Juglans cineria L. 31, 36

Caragana, Caragana arborescens Lam. 19, 29, 30

Catalpa, Northern, Catalpa speciosa Warden. 5, 6, 36

Cherry, Prunus spp. 4, 14, 17, 18, 22, 25, 36, 38, 39

Bird, P. padus L. 18

Bitter, P. emarginata (Douglas) Eaton. 18

Black, P. serotina Ehrh. 14, 18

Canada plum, P. nigra (Aiton). 18

Chokecherry, P. virginiana L. 4, 14, 17, 18, 25

Japanese flowering, Prunus serrulata 14

Mahaleb, P. mahaleb L. 18

Nanking, P. tomentosa Thunb. 4, 18

Pin, P. pensylvanica L. 18

Sandcherry, P. pumila L. 4, 18
212 USDA Forest Service RMRS-GTR-335. 2016.

Sour cherry, P. cerasus L. 4, 14, 17, 18

Sweet cherry, P. avium (L.) L. 4, 14, 18

Western sandcherry, P. besseyi Bailey. 4

Chestnut, Castanea spp. 3

Chokeberry, Aronia spp. 52

Cotoneaster, Cotoneaster spp. 26, 38, 52

Peking, C. acutifolius. 38

Cotinus spp. 36

Cottonwood, Populus spp.1, 9, 10, 22, 23, 24, 28, 29, 32, 38, 39, 40

Eastern, P. deltoides Bartr. 24, 40

Plains, Populus deltoides subsp. monilifera (syn Populus deltoides var. occidentalis)

(Aiton) Eckenw. 1, 9, 24

Crabapple, Malus spp. 5, 8, 17, 22, 26, 36, 52

Currant, Ribes spp. 19, 30

Cypress, Cupressaceae. 5, 42, 47, 50

Arizona, C. arizonica (Green). 42

Dogwood, Cornus spp. 1, 5, 8, 16, 40

Flowering, C. florida L. 5

Pagoda, C. alternifolia L. 1

Douglas-fir, Pseudotsuga (Mirb.) Franco. 48, 53, 55

Elderberry. Sambucus spp. 19

Elm, Ulmus spp. 2, 6, 7, 16, 19, 20, 21, 30, 32, 34, 36, 37, 40

American, U. americana L. 15, 16, 21, 32, 34, 37

Asiatic. U. davidiana var. japonica (Rehder) Nakai. 34

Cedar, U. crassifolia Nutt. 34, 37

Cherry-bark, U. villosa Brandis. 34

Chinese, U. parvifolia Jacq. 34, 37

Evergreen, U. parvifolia Jacq. 34, 37

Japanese, U. japonica (Rehd.) Sarg. 34

Manchurian, U. laciniata (Trautv.) Mayr. 34

Red, U. rubra Muhl. 34, 37

Rock, U. thomasii Sarg. 34

September, U. serotina Sarg. 37

Siberian, U. pumila L. 6, 16, 19, 20, 30, 34, 37

Slippery, U. rubra Muhl. 34, 37

U. rubra × U. pumila hybrid. 37

White, U. americana L. 15, 16, 21, 32, 34, 37

Winged, U. alata Michx. 34, 37

Euonymus, Euonymus spp. 21
USDA Forest Service RMRS-GTR-335. 2016. 213

Fir, Abies spp. 1, 36, 45, 48, 49, 53, 55, 56

Balsam, Abies balsamea (L.) Mill. 55

Douglas, Pseudotsuga menziesii (Mirb.) Franco. 48, 53, 55

White/Concolor, Abies concolor (Gordon & Glend.) Lindl. ex Hildebr. 1

Flowering-quince, Chaenomeles spp. 52

False-cypress, Chamaecyparis spp. 47

Firethorn, Pyracantha spp. 26 

Grape, Vitis spp. 12

Hackberry, Celtis occidentalis L. 8, 15, 33, 36, 40

Hawthorn, Crataegus spp. 5, 8, 26, 36, 52

Hemlock, Tsuga (Endl.) Carr. 39

Hickory, Carya spp. 15, 16, 21, 36

Mockernut, C. tomentosa Nutt. 21

Pecan, C. illinoensis (Wangenh.) K. Koch. 15

Holly, Ilex spp. 5, 13, 16

Honeylocust, Gleditsia spp. L. 1, 6, 19, 21, 30, 36, 39

Native thorny, G. triacanthos var. triacanthos. L. 21

Oriental, G. japonica Miq. 21

Thorny and podless, G. triacanthos var. inermis (L.) C.K. Schneid. 1, 21

Honeysuckle, Lonicera spp. 38

Tatarian, L. tatarica L. 38

Hophornbeam, Ostrya spp. 36

Hornbeam, Carpinus spp. 15, 39

Horsechestnut, Aesculus hippocastanum L. 30, 39

Juneberry, Amelanchier spp. (see also Serviceberry). 5, 17, 25, 26, 36, 52

Juniper, Juniperus spp. 5, 30, 42, 44, 47, 50, 52, 56

Chinese, J. chinensis L. 42, 44, 50, 52

Creeping, J. horizontalis Moench. 42, 47, 52

One-seed, J. monosperma (Engelm.) Sarg. 30

Redcedar (eastern redcedar), J. virginiana L. 30, 42, 44, 47, 50, 52, 56

Rocky mountain, J. scopulorum Sarg. 42, 44, 47, 50, 52, 56

Savin, J. sabina L. 44, 47, 50

Kentucky coffeetree, Gymnocladus dioicus (L.) K. Koch. 36

Larch, Larix spp. 10, 36, 53, 55

Lilac, Syringa spp. 12, 19, 37, 46

Common, S. vulgaris L. 12

Dwarf Korean, S. meyeri C.K. Schneid. 12, 37

Early flowering, S. × hyacinthiflora. 12

Japanese tree, S. reticulata (Blume) H. Hara. 12, 37

Late, S. villosa Vahl. 12, 37
214 USDA Forest Service RMRS-GTR-335. 2016.

Littleleaf, S. pubescens subsp. microphylla Diels. 12

Manchurian/Miss Kim Korean, S. pubescens subsp. patula (Palib.) M.C. Chang &

X.L. Chen. 12, 37

Meyer, S. meyeri C.K. Schneid. 12, 37

Pekin, S. pekinensis Rupr. 12

Preston, S. × prestoniae McKelvey. 12

Linden, Tilia spp. 1, 2, 6, 8, 19, 36

Little leaf, T. cordata Mill. 1, 8

Locust, Robinia spp. 30, 36

Black, R. pseudoacacia L. 30, 36

London plane-tree, Platanus × acerifolia spp. 2

Magnolia, Magnolia spp. 5, 36

Maple, Acer spp. 1, 2, 5, 6, 7, 8, 13, 15, 16, 19, 21, 25, 29, 30, 36, 38, 39

Amur, A. ginnala Maxim. 5, 13

Bigleaf, A. macrophyllum Pursh. 13

Boxelder, A. negundo L. 1, 6, 13, 30, 38

Freeman, A. x freemanii. 5

Hedge, A. campestre L. 1, 13

Miyabe, A. miyabei Maxim. 1

Mountain, A. spicatum Lam. 13

Norway, A. platanoides L. 1, 13, 36

Silver, A. saccharinum L. 5, 13

Striped, A. pensylvanicum L. 13

Sugar, A. saccharum Marsh. 8, 13

Sycamore, A. pseudoplatanus L. 2, 13, 15, 16, 36, 39

Vine, A. circinatum Pursh. 13

Medlar, Mespilus. 52

Mountain-ash, Sorbus spp. 19, 25, 26, 36, 52

Mulberry, Morus spp. 32, 36, 39

Ninebark, Physocarpus spp. 19

Oak, Quercus spp. 1, 2, 3, 5, 6, 7, 8, 12, 21, 30, 35, 36, 40

Black, Q. velutina Lam. 7, 35

Blackjack, Q. marilandica Münchh. 35

Bur, Q. macrocarpa Michx. 1, 3, 5, 6, 7, 8, 21, 30, 35, 36

Q. macrocarpa var. depressa (Nutt.) Engelm. 3

Q. macrocarpa var. macrocarpa Michx. 3

Q. macrocarpa var. oliviformis (Michx. f.) A. Gray. 3

Dwarf chinkapin, Q. prinoides Willd. 35

Golden red oak, Q. rubra ‘Aure.’ 1

Live, Q. virginiana Mill. 30, 35
USDA Forest Service RMRS-GTR-335. 2016. 215

Northern red, Q. rubra L. 1, 2, 5, 6, 7, 8, 15, 35, 36

Pin, Q. palustris Muenchh. 1, 5, 6, 7, 35, 36

Post, Q. stellata Wagenh. 15, 35

Scarlet, Q. coccinea Muenchh. 7

Shumard, Q. shumardii Buckley. 35

Swamp white, Q. bicolor Willd. 1, 5

White, Q. alba L. 7, 8, 35, 36, 40

Peach, Prunus persica Batsch. 4, 5, 7, 14, 17, 18, 25, 30

Pear, Pyrus spp. 17, 26, 36

Persimmon, common, Diospyros virginiana L. 16

Pine, Pinus spp. 1, 5, 6, 36, 38, 40, 41, 43, 46, 48, 51, 53, 54, 55, 56

Austrian, P. nigra Arnold. 1, 43, 44, 46, 51, 55, 56

Eastern white, P. strobus L. 1, 41

Jack, P. banksiana Lamb. 51, 54

Japanese black, P. thunbergiana Franco. 55

Loblolly, P. taeda L. 40, 55

Lodgepole, P. contorta Dougl. 38, 43, 51, 54

Longleaf, P. palustris Mill. 41, 55

Mugo, P. mugo Turra. 1, 41, 43, 46, 51, 54

Ponderosa, P. ponderosa Dougl. ex Laws. 1, 38, 41, 43, 46, 51, 54, 56

Radiata/Monerey, P. radiata D. Don. 46

Red, P. resinosa Ait. 51

Scots/Scotch, P. sylvestris L. 6, 38, 41, 43, 46, 51, 54, 55

Shortleaf, P. echinata Mill. 55

Platycladus orientalis (L.) Franco. 42, 50

Platanus spp. 2, 15, 36, 39

Platanus × acerifolia spp. 2

Platanus occidentalis. 2, 15, 36, 39

Plum, Prunus spp. 4, 7, 14, 17, 18, 22, 30, 38

American, P. americana Marsh. 14, 17, 18, 30

Japanese, P. salicina Lindl. 14

Poplar, Populus spp. 1, 5, 9, 10, 19, 22, 23, 24, 27, 28, 29, 30, 32, 38, 39, 40

Aigeiros, Populus Sect. Aigeiros. 9, 23

Balsam, P. balsamifera L. 9, 23, 24, 29

Tacamahaca Populus sect. Tacamahaca (see Balsam poplar). 23

Bigtooth, P. grandidentata Michx. 27

Black cottonwood, P. trichocarpa Torr. & Gray. 24, 29

Black poplar, P. nigra L. 23, 24, 29

Cottonwood, P. deltoides Bartr. 1, 9, 10, 19, 22, 23, 24, 25, 28, 29, 32, 38, 39, 40

Eastern cottonwood, P. deltoides Bartr. 24, 40
216 USDA Forest Service RMRS-GTR-335. 2016.

Eurasian/European aspen, Populus tremula. L. 27

Japanese poplar, P. maximowiczii Henry. 29

Largetooth, P. grandidentata Michx. 27

Laurel poplar, P. laurifolia Ledeb. 9, 29P. euphratica Olivier 9

Lombardy, P. nigra cv. italica Muenchh. 23, 24

Narrowleaf, P. angustifolia James. 24

Plains cottonwood, Populus deltoides subsp. monilifera (Aiton) Eckenw. (syn 

Populus deltoides var. occidentalis Rydb.) 1, 9, 24

P. alba L. 9

P. davidiana Dode. 9

P. euphratica Olivier. 9

P. euramericana Guinier. 9

P. laurifolia (Ledeb). 9, 29

P. pseudosimonii (Kitag.). 9

P. simonii Carrière. 9

P. tomentosa Carrière. 9

Quaking, P. tremuloides Michx. 9, 10, 21, 23, 24, 27

Pyracantha, Pyracantha spp. 26

Quince, Cydonia spp. 25, 52

Redbud, Cercis canadensis L. 6, 36

Rosaceae. 26, 38, 52

Rose, Rosa spp. 12

Russian-olive, Elaeagnus angustifolia L. 1, 16, 19, 20, 21, 36, 40

Serviceberry, Amelanchier spp. (see also Juneberry). 5, 17, 25, 26, 36, 52

A. arborea (Michx. f.) Fernald. 36

Siberian peashrub, Caragana arborescens Lam. 19, 29, 30

Smoketree, Cotinus spp. 36

Spruce, Picea spp. 1, 6, 36, 39, 45, 48, 49, 53, 55, 56

Black Hills, P. glauca (Moench) Voss. 6, 45, 48, 49, 53

Blue (Colorado blue), P. pungens Engelm. 1, 45, 48, 49, 53, 55, 56

Engelmann, P. engelmannii (Parry) Engelm. 45, 48, 53

Norway, P. abies (L.) Karst. 45, 48, 49, 53, 55

Sitka, P. sitchensis (Bong.) Carrière. 45

White, P. glauca (Moench) Voss. 6, 45, 48, 49, 53, 56

Sumac, Rhus spp. 19

Sweetgum, Liquidambar styraciflua L. 5, 39

Sycamore, Platanus spp. 2, 15, 16, 36, 39

American, P. occidentalis L. 2

Tacamahaca (see Balsam poplar). 23

Tulip tree, Liriodendron tulipifera L. 13, 39

Virginia creeper, Parthenocissus spp. 19
USDA Forest Service RMRS-GTR-335. 2016. 217

Walnut, Juglans spp. 1, 2, 16, 20, 30, 31, 36

Arizona, J. major (Torr.) A. Heller. 31

Black (eastern black), J. nigra L. 1, 20, 30, 31

Butternut, J. cineria L. 31

Japanese, J. ailantifolia Carrière. 31

Little, J. microcarpa Berland. 31

Northern black, J. californica S. Watson. 31

Persian, J. regia L. 31

Southern black, J. hindsii (Jeps.) Jeps. ex R.E. Sm. 31

White-cedar, Chamaecyparis spp. 47

Willow, Salix spp. 1, 9, 10, 13, 19, 21, 22, 25, 30, 32, 36, 38, 39, 40

Golden, S. alba var. vitellina (L.) J. Stokes. 40

Yew, Taxus spp. 5

Index to Plant Diseases and Pathogens

Disease or Pathogen Article Number

Agaricomycetes. 30

Agrobacterium spp. Conn. 22

A. radiobacter (Beijerinck & Van Delden) Conn. 22

A. tumefaciens (Smith & Townsend) Conn. 22

Air pollutant damage. 1

Alcohol flux. 32

Amphiporthe leiphaemia (Fr.) Butin. 2, 3

Anthracnose diseases. 2, 3, 7

Antrodia spp. 30

A. albida (Fr.) Donk. 30

A. juniperina (Murr.) Niem. & Ryv. 30

A. malicola (Berk. & M. A. Curtis) Donk. 30

Apiognomonia spp. 2

A. errabunda (Roberge) Höhn. 2

A. quercina (Kleb.) Höhn. 2

A. veneta (Sacc. & Speg.) Höhn. 2 

Apiosporina morbosa (Schw.) Arx. 18

Armillaria root rot. 38

Armillaria spp. 38

Artist’s conk. 30, 39

Ash yellows. 37

Asteroma spp. 2

Asteroma ulmea (Miles) B. Sutton. 2

Aureobasidium apocryptu (Ellis & Everh.) Herm.-Nijh. 2
218 USDA Forest Service RMRS-GTR-335. 2016.

Basidiomycota. 30

Biscogniauxia canker. 15

Biscogniauxia spp. 15

B. atropunctata (Schwein.) Pouzar. 15

B. mediterranea (De Not) Kuntze. 15

Black knot. 18

Black leaf spot. 2

Black-spot Nectria canker. 19, 20, 21

Blossom blight. 17

Blumeriella jaapii (Rehm.) Arx. 4

Botryosphaeria spp. 3, 42, 44, 50, 51

B. corticola A.J.L. Phillips, A. Alves & J. Luque. 3

B. stevensii Shoemaker. 50, 51

Botryodiplodia canker. 16, 19, 20

Botryodiplodia disease. 16, 20

Botryodiplodia spp. 16, 19, 20

B. hypodermia (Sacc.) Petr. & Syd. 16, 19

B. theobromae Pat. 16, 20

Botryosphaeria rhodina (Berk. and Curt.) v. Arx. 42

B. ulmicola (Ellis & Everh.) Buisman. 16

Brown rot fungi. 17

Brown spot needle blight. 16

‘Candidatus Phytoplasma fraxini’. 37

‘Candidatus Phytoplasma pruni’. 14

‘Candidatus Phytoplasma ulmi’. 37

Cedar apple rust. 42, 44, 50, 52

Ceratocystis fagacearum (Bretz) Hunt. 35

Cercospora blight. 42

Cercospora spp. 3, 4, 11, 42

C. circumscissa Sacc. 3, 4

C. fraxini, (DC.) Sacc. 11

C. sequoiae Ell. & Ev. 42

C. sequoiae var. juniperi Ell. & Ev. 42

Cerrena unicolor (Bull.) Murrill. 30

Cherry leaf spot. 4

Chlorosis. 1, 5, 35, 37, 40

Chlorotic halo. 2

Chondroplea populea (Saccardo & Briard) Klebahn. 23

Coccomyces spp. 3, 4

C. hiemalis Higgins. 3, 4
USDA Forest Service RMRS-GTR-335. 2016. 219

C. lutescens Higgins. 3, 4

Colletotrichum gloeosporioides (Penz.) Penz. & Sacc. 2

Coriolopsis spp. 30

C. floccosa (Jungh.) Ryvarden. 30

C. gallica (Fr.) Ryvarden. 30

Coryneum carpophilum (Lév.) Jauch. 3, 4 

Crown gall. 22

Cryptodiaporthe spp. 23

C. populea (Sacc.) Butin. 23

Cryptosphaeria canker. 24

C. ligniota (Fr.) Auersw. 24

C. populina (Pers.) Sacc. 24

Cyclaneusma needle cast. 46

Cyclaneusma spp. 46

C. minus (Butin) DiCosmo, Peredot & Minter. 46

C. niveum (Pers.) DiCosmo, Peredo & Minter. 46

Cylindrosporium spp. 3, 4, 11

C. padi P. Karst. ex Sacc. 3, 4

C. fraxini (Ellis & Kellerm.) Ellis & Everh. 11

Cytospora canker. 22, 23, 24, 25, 28, 29, 53 

Cytospora spp. 23, 24, 25, 28, 29, 53

C. chrysosperma Pers. ex Fr. 25

C. kunzei Sacc. 53

Cytosporina spp. 24

Daedaleopsis confragosa (Bolton) J. Schröt. 30

Daedalia spp. 30

D. ambigua Berk. 30

D. confragosa Bolt. 30

D. unicolor Bull. 30

Datronia spp. 30

D. scutellata (Schwein.) Gilb. & Ryvarden. 30

D. mollis (Sommerfelt) Donk. 30

Diplodia blight. 51

Diplodia spp. 16, 50, 51

D. gossypina Ellis & Everh. 16

D. mutila (Fr.) Mont. 50

D. natalensis P. Evans. 16

D. pinea (Desm.) J. J. Kickx. 51

D. sapinea (Fr.) Fuckel. 51

D. scrobiculata J. de Wet, Slippers & M.J. Wingf. 51
220 USDA Forest Service RMRS-GTR-335. 2016.

D. theobromae (Pat.) Now. 16

Discosporium populeum (Saccardo) B. Sutton. 23

Discula spp. 2, 3 

D. campestris (Passerini) Arx. 2

D. fraxinea (Peck) Redlin & Stack. 2

D. quercina (Cooke) Sacc. 2, 3

Dothichiza canker. 23, 28

Dothichiza populea Sacc. & Br. 23

Dothistroma spp. 41, 43

D. pini Hulbary. 41, 43

D. septosporum (Dorog.) M. Morelet. 41, 43

Drepanopeziza spp. 9

D. punctiformis Gremmen. 9

D. populi-albae (Kleb.) Nannf. 9

D. populorum (Desm.) Höhn. 9

D. sphaerioides (Pers.) Höhn. 9

Dutch elm disease. 16, 34, 37

Elm leaf blister (see also Taphrina diseases). 7

Elm phloem necrosis. 37

Elm yellows. 37

Endocronartium harknessii (J.P. Moore) Y. Hirats. 54

Entoleuca mammata (Wahlenberg) J.D. Rogers & Y.M. Ju. 27

Environmental stress. 1, 15, 20, 22, 24, 40, 48, 53, 55

Erwinia amylovora (Burrill) Winslow et al. 25, 26

Fire blight. 25, 26

Fomes spp. 30

F. ellisianus Anderson. 30

F. fomentarius (L. ex Fr.) Kickx. 30

F. fraxinophilus (Peck) Sacc. 30

F. ohiensis (Berk.) Murr. 30

Fomitiporia spp. 30

F. punctata (P. Karst.) Murrill. 30

F. texana (Murrill) Nuss. 30

Fomitopsidaceae. 30

Fomitopsis spp. 30

F. cajanderi (Karst.) Kolt. & Pouz. 30

F. melliae (Underw.) Gilb. 30

Fulvifomes robiniae (Murrill) Murrill. 30

Fusarium spp. Link. 28, 29

Fuscoporia gilva (Schwein.) T. Wagner & M. Fisch. 30
USDA Forest Service RMRS-GTR-335. 2016. 221

Ganodermataceae. 30

Ganoderma spp. 15, 30, 39

G. applanatum (Pers.:Wallr.) Pat. 15, 30, 39

G. lobatum (Schw.) Atk. 30 

Geosmithia morbida M. Kolarík, E. Freeland, C. Utley & N.Tisserat. 31

Gloeosporium apocryptum Ell. & Ev. 2

Gloeosporium leaf spot and anthracnose. 2

Gnomonia leaf spot. 2

G. quercina Kleb. 2

Gymnosporangium rusts. 52

Gymnosporangium spp. 52

G. bethelii Kern. 52

G. clavariforme (Jacq.) D.C. 52

G. clavipes C. & P. 52

G. connersii Parmelee. 52

G. corniculans Kern. 52

G. exiguum Kern. 52

G. globosum Farl. 52

G. gracile Pat. 52

G. juniperi-virginianae Schw. 52

G. nelsonii Arth. 52

G. nidus-avis Thaxt. 52

G. trachysorum Kern. 52

G. yamadae Miyabe. 52

Heart rot. 30

Helicotylenchus spp. 40

Herbicide. 1,6, 8, 12, 14, 19, 21, 25

Hoplolaimus spp. Cobb. 40

Hymenochaetaceae. 30

Hypoxylon canker (see also Biscogniauxia canker). 15, 27

Hypoxylon spp. 15, 27 

H. atropunctatum (Schw. ex Fr.) Cke. 15

H. mammatum (Wahl.) Miller. 27

H. mediterraneum (De Not.) J.H. Mill. 15

Inonotus spp. 30

I. dependens (Murrill) Vlasák & Y.C. Dai. 30

I. linteus (Berk. & M.A. Curtis) Teixeira. 30

I. weirianus (Bres.) T. Wagner & M. Fisch. 30

Isthmiella crepidiformis (Darker) Darker. 45

Kabatina juniperi Schneider & v. Arx. 44
222 USDA Forest Service RMRS-GTR-335. 2016.

Kabatina tip blight. 42, 44, 48

Laetiporus sulphureus (Bulliard) Murr. 30

Lasiodiplodia theobromae (Pat.) Griffon & Maubl. 16

Leaf blister. 6, 7 

Leaf lesion. 3, 4, 9, 11, 13, 44

Leaf rust. 10

Leaf scorch. 1

Leaf spot. 2, 3, 4, 7, 9, 11, 29

Leaf tatters. 8

Lecanosticta acicola (Thüm.) Syd. 41

Lepteutypa cupressi (Nattrass, C. Booth & B. Sutton) H.J. Swart. 50

Leucocytospora kunzei (Sacc.) Z. Urb. 53

Leucostoma spp. 25, 53

L. kunzei (Fr.) Munk. 25, 53

Libertella spp. 24

Lilac witches’-broom. 37

Lirula needle blight. 45

Lirula spp. 45

L. macrospora (Hartig) Darker. 45

L. brevispora Ziller. 45

Lophodermium piceae (Schrad.) Chevall. 45

Maple leaf curl (see Taphrina diseases). 7

Maple tar spot. 2

Marssoniella juglandis (Lib.) Höhn. 2 

Marssonina leaf spot. 9

Marssonina spp. 9, 11

M. balsamiferae Y. Hirats. 9

M. brunnea (Ell. & Ev.) Magnus. 9

M. brunnea f. sp. brunnea Spiers. 9

M. brunnea f. sp. trepidae Spiers. 9

M. castagnei (Desm. & Mont.) Magnus. 9

M. fraxini Ell. & Davis. 11

M. populi (Lib.) Magn. 9

M. salicicola (Bres.) Magnus. 9

Melampsora rust. 9, 10

Melampsora spp. 9, 10, 29

Melampsora × Columbiana. 10

M. epitea Thüm. 10

M. medusae Thuem. 10

M. occidentalis H.S. Jacks. 10
USDA Forest Service RMRS-GTR-335. 2016. 223

Meloidogyne spp. 40

Monilinia spp. 17

M. amelanchieris (J.M. Reade) Honey. 17

M. demissa (B.F. Dana) Honey. 17

M. fructicola (Wint.) Honey. 17

M. laxa (Aderh. & Ruhland) Honey. 17

Mycosphaerella leaf spot. 11

Mycosphaerella spp. 4, 11, 29, 41

M. cerasella Aderh. 4

M. dearnessii M.E. Barr. 41

M. effigurata (Schw.) House. 11

M. fraxinicola (Schw.) House. 11

M. jaczewskii Potebnia. 29

M. pini Rostr. 41, 43

M. populi (Auersw.) J. Schröt. 29

M. populorum G.E. Thomps. 29

Naemacyclus minor Bulin. 46

Nectria spp. 19, 20, 21

N. cinnabarina Tode ex Fr. 19, 20, 21

N. nigrescens Cooke. 19, 20, 21

Oak wilt. 3, 35, 

Oakleaf blister (see Taphrina diseases). 7

Ophiostoma spp. 34

O. novo-ulmi Brasier. 34

O. ulmi (Buisman) Melin & Nannf. 34

Oxyporus populinus (Schum.) Donk. 30

Passalora spp. 4, 42

P. circumscissa (Sacc.) U. Braun. 4

P. sequoia (Ellis & Everh.) Y.L. Guo & W.H. Hsieh. 42

Peach leaf curl (see also Taphrina diseases). 7

Perenniporia spp. 30

P. ellisiana (F.W. Anderson) Gilb. & Ryvarden. 30

P. fraxinophila (Peck) Ryvarden. 30

P. ohiensis (Berk.) Ryvarden. 30

Peridermium spp. 54

P. harknessii J.P. Moore. 54

Phellinopsis conchata. 30

Phellinus spp. 30

P. alni. 30

P. conchatus (Pers. ex Fr.) Quel. 30
224 USDA Forest Service RMRS-GTR-335. 2016.

P. everhartii (Ell. & Gall.) Ames. 30

P. gilvus (Schw.) Pat. 30

P. igniarius (L. ex Fr.) Quel. 30

P. pomaceus (Pers. ex S.F. Gray) Maire. 30

P. punctatus (Fr.) Pilat. 30

P. ribis (Schum. ex Fr.) Quel. 30

P. robineae (Murr.) A. Ames. 30

P. robustus (P. Karst.) Bourdot & Galzin. 30

P. tremulae (Bond.) Bond. & Boriss. 30

P. tuberculosus (Bawmg) Niemala. 30

P. weirianus (Bres.) Gilbn. 30

Phellinus stem decays. 30

Phomopsis blight. 42, 47, 48

Phomopsis canker. 20, 28 

Phomopsis spp. 20, 22, 45, 47, 48, 49

P. arnoldiae Sutton. 20

P. elaeagni (Carter & Sacamano) Arnold & Carter. 20

P. juniperovora Hahn. 44, 47

Phyllactinia fraxini (D.C.) Fuss. 2

Phyllosticta leaf spot. 11

Phytoplasma. 14, 37

Piggotia fraxini Berk. & M.A. Curtis. 11

Pine wilt disease. 41, 55

Plagiostoma fraxini (Redlin & Stack) Sogonov. 2

Pleonectria spp. 21.

P. austroamericana Speg. 19, 21

Plum pockets (see Taphrina diseases). 7

Podosphaera phytoptophila (Kellerm. & Swingle) U. Braun & S. Takam. 33

Polyporaceae. 30

Polypores. 30

Powdery mildew. 12, 33

Pseudocercospora juniperi (Ellis & Everh.) B. Sutton & Hodges. 42

P. juniperi (Ellis & Everh.) B. Sutton & Hodges. 42

P. fraxini (Ellis & Kellerm.) U. Braun. 11

Pseudomonas spp. Migula. 4

P. syringae pv. syringae (Van Hall) Dye. 4

Quince rust. 52

Red band disease. 43

Rhizosphaera kalkhoffii Bub. 48

Rhizobium radiobacter. 22

Rhizosphaera needle cast. 48, 49
USDA Forest Service RMRS-GTR-335. 2016. 225

Rhytisma spp. 13

R. acerinum Pers. ex Fr. 13

R. americanum Hudler & Banik. 13

R. punctatum (Pers.) Fr. 13

Root lesion. 40, 56

Root-lesion nematodes. 40, 56

Root rot. 3, 30, 38, 39, 56

Rust fungi. 10, 52, 54

Scirrhia acicola (Dearn.) Siggers. 41, 43

Schirria pini (Funk & Parker). 41, 43

Sclerophoma pithyophila (Corda) Höhn. 45

Seiridium unicorne (Cooke & Ellis) B. Sutton. 50, 51

Septoria canker. 29

Septoria leaf spots. 11, 29

Septoria spp. 11, 29

S. aceris (Lib.) Berk. & Br. 29

S. fraxinicola U. Braun. 11

S. musiva Peck. 29

Slime flux. 32

Sphaeropsis spp. Leveille. 16, 51

S. hypodermia Höhn. 16

S. sapinea (Fr.) Dyke & Sutton. 51

S. ulmicola Ellis & Everh. 16

Sphaerulina spp. 29

S. aceris (Libert) Verkley, Quaedvlieg & Crous. 29

S. musiva (Peck) Quaedvlieg, Verkley & Crous. 29

S. populicola (Peck) Quaedvlieg, Verkley & Crous. 29

Stegophora ulmea (Schw.:Fr.) Syd. & Syd. 2

Stem lesion. 29, 40

Stigmina needle cast. 48, 49

Stigmina spp. 48, 49

S. carpophila (Lév.) M.B. Ellis. 4

S. lautii B. Sutton. 45, 49

Streptomyces avermitilis. 55

Stress. 1, 2, 3, 5, 7, 8, 10, 12, 13, 14, 15, 16, 19, 20, 21, 22, 23, 25, 28, 32, 36, 37, 38, 40, 

46, 48, 50, 51, 53, 55

Sycamore anthracnose. 2

Sydowia polyspora (Bref. & Tavel) E. Müll. 45

Sydowiellaceae. 2

Taphrina diseases. 7

Taphrina spp. 7
226 USDA Forest Service RMRS-GTR-335. 2016.

T. caerulescens (Mont. & Desm.) Tul. 7

T. carveri Jenkins. 7

T. communis (Sadeb.) Grs. 7

T. deformans (Berk.) Tul. 7

Tar spot. 13

Thyronectria canker. 21

Thyronectria austroamericana (Speg.) Seeler. 21

Thyrostroma carpophilum (Lév.) B. Sutton. 4

Trametes spp. 30

T. elegans (Spreng.) Fr. 30

T. hispida Bagl. 30

T. malicola Berk. & Curt. 30

T. mollis (Sommert.) Fr. 30

T. rigida Berk. & Mont. 30

T. sepium Berk. 30

T. suaveolens (L.:Fr.) Fr. 30

T. trogii Berk. 30

Trametes stem decay. 30

Trichothecium roseum (Pers.) Link. 18

Tubakia spp. 3, 35

T. iowensis T.C. Harr. & D. McNew. 3, 35

Tubercularia spp. 19, 20, 21

T. ulmea Carter. 16, 19, 20, 21

T. vulgaris Tode: Fr. 19, 21

Tubercularia canker. 19, 20, 21

Tubercularia spp. 19, 20, 21

T. ulmea Carter. 16, 19, 20, 21

T. vulgaris Tode: Fr. 19, 21

Twig blight. 2, 3, 44, 45

Valsa spp. 25, 53

V. kunzei (Fr.) Fr. 53

V. sorida Nits. 25

Valsaceae. 2

Verticillium spp. Nees. 36

V. albo-atrum Reinke & Berth. 36

V. dahliae Kleb. 36

Verticillium wilt. 36

Viruses. 14, 33, 40

Virus-X. 14

Walnut anthracnose. 2
USDA Forest Service RMRS-GTR-335. 2016. 227

Western gall rust. 54 

Western X-disease. 14

Wetwood. 32

White-mottled heart rot. 39 

Wilsonomyces carpophilus (Lév.) Adask., J.M. Ogawa & E.E. Butler. 4

Witches’-broom. 1, 33, 37, 52, 53, 54

Xanthomonas spp. Dowson. 4

X. arbicola pv. pruni. 4

X-disease. 14

Yellows diseases. 37

Index to Associated Insects, Nematodes, and 

Pathogen Vector

Insect Article Number

Beetles. 3, 14, 15, 17, 31, 34, 35, 37, 55

Banded elm bark beetle, Scolytus schevyrewi Semenov. 34

Bark beetle. 14, 34, 37

Carpophilus sayi Parsons. 35

Emerald ash borer, Agrilus planipennis. 35

Monochamus spp. 55 

Carolina pine sawyer beetle, M. carolinensis (Oiiv.). 55

Pine sawyer beetle, M. alternalus (Hope). 55

Southern pine sawyer beetle, M. titillator (F.). 55

White-spotted pine sawyer, M. scutellatus (Say). 55

Native elm bark beetle, Hylurgopinus rufipes Eichh. 34 

Plum curculio, Conolrachelus nenuphar (Herbst.). 17

Sap beetles (Coleoptera: Nitidulidae). 35

Smaller European elm bark beetle, Scolytus multistriatus (Marsham). 34

Two-lined chestnut borer, Agrilus bilineatus Weber. 3, 15

Walnut twig beetles, Pityophthorus juglandis Blackman. 31

Eriophyid mite, Aceria celtis, syn. Eriophyes celtis. 33

Fruit flies, Drosophila spp. 17

Leafhoppers. 14, 37 

Cicadellidae. 37

Latalus spp. 37

Scaphoideus spp. 37

Midges. 43

Juniper midge, Contarinia spp. 43

Moths. 17, 43

Juniper webworm, Dichomeris sp. 43
228 USDA Forest Service RMRS-GTR-335. 2016.

Oriental fruit moth, Grapholita molesta Busck. 17

Nematodes. 40, 55, 56

Dagger nematode, Xiphinema americanum Cobb. 40, 56

Lance nematode, Hoplolaimus spp. Cobb. 40

Pin nematode, Paratylenchus spp. 40, 56

P. penetrans (Cobb) Filipjev & Stekhoven. 56

Pinewood nematode, Bursaphelenchus xylophilus (Steiner & Buhrer) Nickle. 55

Root-knot nematode, Meloidogyne spp. 40

Root-lesion nematode, Pratylenchus spp. 40

Spiral nematode, Helicotylenchus spp. 40

Stunt nematode, Tylenchorhynchus spp. 40, 56

T. ewingi Hooper. 40, 56

Planthoppers. 37 

Spittlebugs. 37 

Lepyronia quadrangularis Say. 37

Index to Associated Associated Chemicals

Chemical Article Number

2,4-D / 2,4-D-DP. 6

Acetochlor. 8

Atrazine. 6

Azoxystrobin. 45, 48, 49

Benzoic acids. 6

Benzothiadiazoles. 6

Bromacil. 6

Bromoxynil. 6

Carbamates. 6

Chlorothalonil. 18, 46, 48, 49

Clopyralid. 6

Contact herbicides. 6

Copper hydroxide. 18, 45, 48, 49

Copper oxychloride. 48

Copper salts of fatty and resin acids. 45, 48, 49

Copper sulfate. 45, 48, 49

Desmedipham. 6

Dicamba. 6

Dimethenamid. 8

Fluroxypyr. 6

Glyphosate. 6

Herbicide damage. 1, 6
USDA Forest Service RMRS-GTR-335. 2016. 229

Hormonal herbicides. 1

Mancozeb. 18, 45, 48, 49

MCPA, MCPP. 6

Metolachlor. 8

Mono- and di-potassium salts of phosphoric acid. 45, 48, 49

Nitriles. 6

Non-hormonal type herbicides. 1 

Phenoxy herbicides. 6

Phenoxyacetic acids. 6

Phenylureas. 6

Picloram. 6

Propiconazole. 3, 48, 49

Pyraclostrobin. 45, 48, 49

Pyridine carboxylic acids. 6

Simazine. 6

Sulfur. 4, 45, 48, 49

Tebuthiuron. 6

Terbacil. 6

Thiophanate-methyl. 18, 45, 48, 49

Triazines. 6

Triclopyr. 6

Trifloxystrobin. 48, 49

Uracils. 6

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