Forest Nursery

Diseases

in the

United States C::J

J

Agriculture Handbook No. 470

Forest Service

U.S. Department of Agriculture
Forest Nursery Diseases

in the

United States

Technical Coordinators:

Glenn W. Peterson

Rocky Mountain Forest and Range Experiment Station

Lincoln, Nebraska

and

Richard S. Smith, Jr.

Pacific Southwest Forest and Range Experiment Station

Berkeley, California

Agriculture Handbook No. 470

March 1975

Forest Service

U.S. Department of Agriculture

Library of Congress Catalog No. 74-600103

For sale by the Superintendent of Documents, U.S. Government Printing OfRce

Washington, D.C. 20402 Price $3.50 (paper cover)

Stock Number 0100-03299
This handbook was prepared at the request of the North American Forestry Commission's Working Party on Forest Insects and

Diseases. Glenn W. Peterson and Richard S. Smith, Jr., pathologists of the Rocky Mountain and Pacific Southwest Forest and

Range Experiment Stations, respectively, wrote the introductory

sections and authored or coauthored 12 of the 33 individual disease

sections. They also worked over a 3-year period with the other 16

contributing authors to ensure technical accuracy, and uniformity

and thoroughness of treatment.

POLI-OW THE LABEI.
CONTENTS

Page

INTRODUCTION 1

Scope 2

Disease Diagnosis 2

Disease Control 3

Use and Safe Handling of Pesticides 4

I. ROOT AND SOILBORNE DISEASES 5

1. Damping-off 6

T. H. Filer, Jr., and Glenn W, Peterson

2. Fusarium Root Disease 9

Richard S. Smith, Jr,

3. Charcoal Root Disease 11

Richard S. Smith, Jr.

4. Black Root Rot of Pine 14

Charles S. Hodges

5. Phytophthora Root Rot 17

E. G. Kuhlman and Richard S. Smith, Jr.

6. Phytophthora Root Rot of Black Walnut Seedlings 19

Ralph J. Green, Jr.

7. Cylindrocladium Root Rot 23

Charles E. Cordell and Darroil D. Shilling

8. Crown Gall 27

Glenn W. Peterson and Larry W. Moore

9. Nematodes 31

John L. Ruehle

10. Mycorrhizae of Forest Nursery Seedlings 35

Donald H. Marx

II. STEM AND BRANCH DISEASES (CANKERS) 41

11. Phomopsis Canker of Douglas-Fir 42

Richard S. Smith, Jr.

12. Sirococcus Tip Blight 45

Richard S. Smith, Jr.

13. Grey Mold of Giant Sequoia _ 47

Richard S. Smith, Jr.

14. Scleroderris Canker 50

Darroll D. Shilling

15. Southern Fusiform Rust 53

S. J. Rowan

16. Western Gall Rust 56

Richard S. Smith, Jr.

17. Jack Pine Rusts 59

Darroil D. Shilling

18. White Pine Blister Rust 62

Darroil D. Shilling

III. FOLIAGE DISEASES 66

19. Lophodermium Needle Cast of Pines 67

Darroll D. Shilling

20. Brown Spot Needle Blight 69

Albert G. Kais

21. Dothistroma Needle Blight of Pines 72

Glenn W. Peterson

111
Page

22. Phomopsis Blight of Junipers 76

Glenn W. Peterson

23. Cercospora Blight of Juniper _- 80

Charles S, Hodges

24. Rosellinia Needle Blight 83

Richard S. Smithy Jr.

25. Snow Blight of Conifers 85

Darroll D, Shilling

26. Sycamore Anthracnose 88

Winand K. Hock and Frederick H. Berry

27. Walnut Anthracnose 92

Frederick H, Berry and Winand K. Hock

28. Marssonina Lreaf Blight of Bigtooth Aspen 95

Leon S. Dochinger

29. Melampsora Rust on Cottonwood 99

T. H. Filer, Jr,

30. Septoria Leaf Spot and Canker on Cottonwood 101

T. H. Filer, Jr,

31. Yellows or Chlorosis 104

S, J. Rowan

32. Air Pollution 107

Leon S, Dochinger and Keith F. Jensen

IV. STORAGE MOLDS 114

Drake Hocking

GLOSSARY 116

INDEX TO HOST TREES, WITH SCIENTIFIC EQUIVALENTS _ 121

INDEX TO DISEASES 123

U.S. Department of Agriculture, Forest Service.

1975. Forest Nursery Diseases in the United States.

U.S. Dep. Agrie, Agrie. Handb. 470, 125 p.

Discusses hosts, symptoms, damage, life cycles, distribution of

pathogens, and methods for control for 31 diseases of seedlings in

forest nurseries—9, root ; 8, stem and branch ; 14, foliage. Includes

a short discussion on storage molds, plus selected references,

glossary, disease index, and host index.

OXFORD : 443.2232.327.2.

KEYWORDS : seedling diseases, seedling disease control

(nursery.)

The use of trade and company names is for the benefit of the

reader; such use does not constitute an official endorsement or

approval of any service or product by the U.S. Department of

Agriculture to the exclusion of others that may be suitable.
Forest Nursery Diseases

in the

United States

INTRODUCTION

Forest nurseries and the seedlings they grow play an important

part in keeping this Nation's forest lands productive. The need

for more trees and forests, for a host of purposes, is steadily

increasing. Currently, millions of acres of commercial forest

land under private, State, or Federal ownership are nonstocked

or poorly stocked. More of these nonproductive acres are created

each year by fire, pests, and timber harvests. Planting of these

understocked forest lands is one means of avoiding or reducing

a future timber shortage. At the 1972 National Tree Planting

Conference, John McGuire, Chief of the USDA Forest Service,

said, **I believe that tree planting should be the main thrust in

meeting the challenge of providing for American forests.'' Our

forest tree nurseries must play the key role in this program by

providing forest managers with the needed quality and quantity

of planting stock.

Factors that influence the quantity and quality of nursery

seedlings may also influence our future timber supply. These

factors include seedling diseases caused by fungi, bacteria, and

nematodes. These diseases, which are active from the time of

sowing through outplanting, may kill seedlings directly, or stunt

or malform them so they must be rejected. Nursery diseases may

also lower field survival of outplanted seedlings. Lastly, nursery

diseases may be a threat to our forests when infected seedlings

are planted in forested areas where the disease does not and has

not existed. The introduction of white pine blister rust into North

America on seedlings is the most notable example of this kind

of disaster.

The economic losses due to diseases in forest nurseries must

include more than just the cost of producing the dead and culled

seedlings. They include the cost of a second site preparation of

the plantation when plantable seedlings are not available due to

disease losses at the nursery, or the cost of a second planting or

interplanting when seedling diseases continue to cause mortality

in the plantation. The basic land cost must also be included when

forest land is held out of production by nursery disease problems.

Thus the economic losses resulting from nursery diseases are

not restricted to the nursery operation; impacts on the forests

may be even greater.
SCOPE

The purpose of this Handbook is to assist the nurseryman,

forester, extension pathologist, and others in identifying, evaluating, and controlling the diseases encountered in forest nurseries.

In preparing this Handbook we have tried to bring together

all the currently available information on seedling diseases that

have been a problem in our forest tree nurseries. In most cases,

each disease is treated separately in the following standard

format: Distribution, Hosts and Damage, Life History, Control,

and Selected References. This information has been gathered

and presented by authorities who in most cases have worked

with these individual disease problems in the field. Limited

coverage of some diseases indicates our lack of knowledge about

them. General information on disease diagnosis, disease control,

and pesticide handling is included in the introduction. A glossary,

a disease index, and a host index are in the back of the Handbook.

We have not attempted to cover the subjects of nutrient deficiency, herbicide injury, or environmental injury (except air

pollution). These large subjects could not be adequately treated

in this Handbook, and probably deserve separate coverage. There

are also numerous minor diseases, such as some of the foliage

diseases of hardwoods, about which we have such limited information that we made no attempt to cover them.

If further information or assistance is required, the nurseryman should contact a plant pathologist from one of the agencies

listed below in his area:

Forest Pest Control, Forest Service, U.S. Department of

Agriculture

Agricultural Extension Service, State university

State Department of Agriculture

State Department of Forestry

DISEASE DIAGNOSIS

Within each disease writeup, the sections on distribution, hosts,

and damage will be the most helpful in diagnosing a particular

disease. This diagnosis is an investigation to determine the cause

of a condition or problem. It involves the art of identifying a

disease from its symptoms, signs, and patterns. As in all investigations, it requires a systematic search for clues, and the

assembly of these clues into a reasonable explanation of the

situation. The symptoms (expressions of the diseased host), signs

(evidences of the cause), and patterns of occurrence, are the

clues upon which we must base our determinations.

The following guide may be helpful in diagnosing tree disease :

1. Determine as accurately as possible the part of the plant

which is actually affected. The death of only the needles indicates

a needle disease ; of the stem and/or branches—a canker disease ;

of the whole tree—a root disease, or drought. Note the pattern
of the disease in the seedlings. Is the damage limited to the south

side? to the lower or upper crown?

2. Note what species are affected. Are there any individuals

which are affected less or which are free of the problem?

3. Note the pattern of occurrence. What areas show the

problem most severely? How do these areas differ from those

areas free of the problem? Are these problem areas in any

particular portion of the bed or related to a particular cultural

activity?

4. If the cause of the disease is not immediately evident, look

first for the simplest causes, such as animal damage, frost, simple

injuries, or other obvious causes of trouble.

5. Look for the presence of fungi, insects, or other parasites.

Observe accurately, and try to judge whether the organisms

found are the main cause of the trouble or just secondary.

6. If the whole tree is dead or suffering and nothing is found

aboveground to indicate the cause of the disease, expose the

roots and root crown for examination.

7. If still unsure, learn about the recent history of the problem

and the area. Is the problem of recent origin? When was it first

noted? What cultural practices have been used in the area, such

as the use of herbicides, fertilizers, irrigation, or flooding? Can

these be related through pattern or time of appearance to the

injuries?

DISEASE CONTROL

There are two general types of disease control available to

the nurseryman: cultural and chemical.

In cultural control methods, the nurseryman maintains control

over a disease problem by performing routine nursery operations

in certain prescribed ways. Such prescriptions may dictate specific seedbed densities, time of planting, watering schedule, and

so forth. Most of these cultural methods control a disease problem

by modifying the environment so that it becomes unfavorable for

the disease or its causal organism.

Chemical control methods rely mainly on treatment before the

disease becomes established, either by eradicating the pest from

the area (soil fumigation) or by protecting the plant with a

chemical barrier (foliage sprays). Once a plant has been infected

it is very difficult to cure the plant with chemicals. This means

that the nurseryman must be prepared to apply his chemical

controls either just before or at the first appearance of the

disease.

In some cases the information in this Handbook on control

measures—particularly on fungicides—is very generalized, because pesticide regulations prohibit the recommendation of nonregistered fungicides for plant disease control. As information

required for pesticide registration becomes available, more specific

recommendations can be made.
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 Federal Environmental Protection Agency, consult your county agricultural agent

or State extension specialist to be sure the intended use is still

registered.

The follov^ing rules 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. If used properly, they will not cause injury. The dangers

associated with mishandling and misapplication of pesticides,

however, include possible injury to the operator and handler, and

damage to the seedling crop, 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 contwct.—Avoid inhaling sprays and dusts. Avoid

contact with skin and eyes. When directed by the label, wear the

proper protective clothing and a mask. Do not eat or chew 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 or 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 in doubt on proper

disposal procedures, contact the nearest agricultural 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 get the

patient to a hospital immediately.
I. ROOT AND SOILBORNE DISEASES

Root and soilborne diseases are those diseases of the belowground portions of seedlings which may occur from the time of

planting until lifting. Included in this group are pre- and postemergence damping-off, and diseases of the root crown, taproot,

lateral roots, and tips. The diseases in this group are caused by

bacteria, fungi, and nematodes. A section on mycorrhizae also

is included in this group because they involve an infection of the

root tips, even though beneficial to the host.

Root diseases are some of the most difficult disease problems

to diagnose. The above-ground symptoms of most root diseases

are very similar to one another and also to symptoms caused by

unfavorable soil conditions such as high plow pan, excessive soil

moisture, soil toxicity, and some nutrient déficiences.

The following diseases are considered to be the most important

and common root and soilborne diseases found in North American

forest tree nurseries.
1. DAMPING-OFF

T. H. FILER, JR. AND GLENN W. PETERSON*

Damping-off is one of the first diseases the nurseryman encounters in his newly sown beds. Many fungi cause damping-off,

but those most commonly encountered are Phytophthora spp.,

Pythium spp., Fusarium spp., and Rhizoctonia solani. The disease

sometimes causes 15 percent or greater mortality of the viable

seed, and must therefore be considered of major importance.

Only very young seedlings are attacked ; after the stems begin

to develop woody tissue—perhaps in 4 to 6 weeks after seed

germination—susceptibility declines rapidly.

Distribution and Hosts

Damping-off is widespread in nurseries throughout North

America. Most conifer species are susceptible, with the notable

exception of the junipers. Among the hardwoods, American and

Siberian elms, black locust, Russian mulberry, sweetgum, sycamore, yellow-poplar, and Russian-olive are very susceptible. Green

ash, northern catalpa, hackberry, honeylocust, and bur oak have

considerable resistance.

Symptoms

Symptoms vary, depending on when infection occurs. Early

attacks on the developing radicle may kill the seedling before it

emerges from the soil. Such preemergence damping-off may go

undetected or be explained away as ''poor seed."

Postemergence infection characteristically occurs at or just

below the groundline, resulting in a water-soaked or necrotic area

on the succulent stem. In conifers this tissue collapses, and the

seedling becomes flaccid and topples over. In contrast, hardwood

seedlings usually remain upright, gradually wilt, and break off.

Damping-off is often confused with heat lesions, which appear as

whitish sunken areas on the stem just above the groundline. Heat

lesions, however, usually occur on only one side of the stem.

*Respectively, Principal Plant Pathologist, Southern Hardwoods Laboratory, Southern Forest Experiment Station, USDA Forest Service, Stoneville,

Miss., and Principal Plant Pathologist, Rocky Mountain Forest and Range

Experiment Station, USDA Forest Service, Lincoln, Nebr.
Disease Development

The causal fungi as a rule are native to the nursery soils. Some

live on dead organic material, while others survive as resting or

dormant spores while awaiting a new host to infect. Populations

of these pathogenic fungi tend to build up in nursery soils with

each successive year of cropping, and average seedling losses

tend to increase.

Any condition that reduces seedling growth and vigor usually

causes an increase in infection. The disease usually is most

severe in nursery soils that are excessively wet. A stand of apparently healthy seedlings may show severe damping-oif after

just 1 or 2 rainy days. Sideboards and shading frames that

restrict airflow may increase infection, since the beds will not

dry rapidly. Temperature also influences development of this

disease, depending upon the fungus species involved. Some fungi,

such as Pythium spp., are favored by high temperatures (27°-

35°C). Nitrogen fertilizers applied before or during the period of

seedling susceptibility may also increase losses. Soil pH is significant; losses are usually greatest at high pH's and minimal near

pH 5.5.

Control

Damping-oflf may be controlled by cultural or by chemical

means. Losses have generally been reduced by practices that

increase the rate and uniformity of germination and the speed

at which the seedlings grow in early weeks. Regulation of soil

moisture may reduce losses from certain fungi.

The seed-protectant fungicide thiram is sometimes effective,

but also often inconsistent, in reducing losses in conifer beds.

Fungicides may reduce the amount and speed of germination if

temperatures are high.

Soil treatment prior to sowing is feasible. Applying inorganic

acids to reduce pH has been useful on soils with pH range 6-8.

Drenching soil with the fungicide captan is also eifective, but

once symptoms become evident it is usually too late to drench.

Good results may be achieved by soil fumigation prior to

sowing. Methyl bromide, chloropicrin, vorlex, or combinations

of these and other fumigants give excellent control of dampingoff and other soilborne diseases in both hardwood and pine

seedlings.

Selected References

Filer, T. H., Jr.

1967. Damping-off of sweetgum by Pythium sylvaticum. Phytopathology

57:1284.

Filer, T. H., Jr.

1969. Controlling infectious diseases in nurseries. Southeast. Area,

For. Nurserymen Conf. Proc. 1968:144-148.
Filer, T. H., Jr., and E. R. Toole.

1968. Effects of methyl bromide on mycorrhizae and growth of sweetgum seedlings. Plant Dis. Rep. 52:483-485.

Garrett, S. D.

1956. Biology of the root-infecting fungi. 292 p. Cambridge Univ. Press,

London.

Hodges, Charles S., Jr.

1962. Diseases in southeastern forest nurseries and their control. U.S.

Dep. Agrie. For. Serv. Southeast. For. Exp. Stn., Stn. Pap. 142,

16 p. Asheville, N.C.

Leach, L. D.

1947. Growth rates of host and pathogen as factors determining the

severity of preemergence damping-off. J. Agrie. Res. 75:161-179.

Peterson, Glenn W.

1970. Response of ponderosa pine seedlings to soil fumigants. Plant

Dis. Rep. 54:572-575.

Peterson, Glenn W.

1970. Seed-protectant chemicals affect germination of ponderosa pine

seed. Tree Planters Notes 21(4):25-29.

Rowan, S. J., T. H. Filer, and W. R. Phelps.

1972. Nursery diseases of southern hardwoods. U.S. Dep. Agrie. For.

Pest Leafl. 137, 7 p.

Ruehle, John L., Jack T. May, and S. J. Rowan.

1966. Nursery fumigation trial with vorlex. Tree Planters* Notes 76,

p. 4-7.

Wright, Ernest.

1944. Damping-off in broadleaf nurseries of the Great Plains region.

J. Agrie. Res. 69:77-94.

Wycoff, H. B.

1952. Methyl bromide controls soil organisms which cause mortality of

eastern white pine seedlings. Tree Planters' Notes 12, p. 11-14.
2. FUSARIUM ROOT DISEASE

Fusarium oxysporum Schlect. emend, Snyd. & Hans.

RICHARD S. SMITH, JR.*

Fusarium root disease is one of the most common diseases of

conifer seedlings in the world. Besides causing a root disease,

F. oxysporum and other species of Fusarium are often responsible

for damping-off in earlier stages of seedling development. This

section is concerned only with the root disease. The subject of

damping-off is covered in article 1 of this Handbook.

Distribution

This disease is common and widespread through most of North

America. It has been especially important in the southeastern,

southern, and western parts of the United States. Although it

appears to be favored by the warmer climates, it is not restricted

to these areas.

Hosts and Damage

Most conifer seedlings are susceptible to the Fusarium root

disease. The pathogen is known to exist in several forms which

are host specific. So far as conifer seedlings are concerned, only

one specialized form, F. oxysporum f. sp. pini, has been described.

F. oxysporum attacks and kills the roots of a seedling, causing

chlorosis, stunting, and wilting of the top and eventually death.

As with many root diseases, the eifects are chiefly (a) seedling

mortality in the nursery bed; (b) increased number of cull

(stunted) seedlings; and (c) increased losses after outplanting

because of impaired root systems.

Life History

The pathogenic forms of F, oxysporum, like so many other root

disease fungi, are inactive in the soil in the absence of a host.

They usually remain dormant in the form of chlamydospores—

microscopic, thick-walled, single-celled resting spores. When a

seedling root grows by a dormant chlamydospore, exudates from

the root supply nutrients to the spore and stimulate its germination. The fungus grows over the root surface, penetrates

between two epidermal cells, and spreads intracellularly through

^Principal Plant Pathologist, Pacific Southwest Forest and Range Experiment Station, USDA Forest Service, Berkeley, Calif.

9
the cortex. The fungus colonizes the cortex and the xylem elements of the infected seedling. The exact cause of seedling

death is not clear. Most pathogenic forms of F. oxysporum cause

a vascular wilt, although many descriptions of the disease in

conifers suggest it is a cortical rot. Either or both may be

involved in seedling death.

After the pathogen has killed the seedling and has colonized

the host tissue, it converts much of its mass into macroconidia

and chlamydospores. The macroconidia are large, multicelled

spores which may infect other roots, but which usually convert

themselves into chlamydospores—when conditions become unfavorable—and await next season's crop.

The Fusarium fungus may be carried with the plant when it is

outplanted and continue to attack the infected seedling roots,

causing unexpectedly high transplant mortality. Although this

fungus may continue activity for a year or two, the outplanted

seedling may outgrow the infection if the infection is light.

Studies in the West have shown that Fiisarmm disappears after

3 or 4 years from the roots of seedlings outplanted into the

forest. Thus, although the disease may be the cause of first-year

transplant mortality, it will not persist in the forest environment

and cause continuing problems.

Control

Cultural methods on the whole have not been effective in controlling this disease. Soil amendments, which showed promise in

controlling certain Fusarium diseases of agricultural crops, have

not yet proved to be adequately effective in the forest nursery.

Fumigating the soil with methyl bromide, chloropicrin, or other

soil fumigants before planting currently is the best method of

control. Fumigation has been used effectively in many parts of

North America. Soil fungicides have in most cases been less

effective than soil fumigants.

Selected References

Bloomberg, W. J.

1966. The occurrence of endophytic fungi in Douglas-fir seedlings and

seed. Can. J. Bot. 44:413-420.

Hodges, Charles S.

1962. Black root rot of pine seedlings. Phytopathology 52:210-219.

Matuo, Takken, and Osamu Chiba.

1966. Species and formae spéciales of Fusaria causing damping-off and

root-rot of coniferous seedling in Japan. Ann. Phytopath. Soc.

Japan 32:14-22.

Smith, Richard S., Jr.

1967. Decline of Fusarium oxysporum in the roots of Pinus lambertiana

seedlings transplanted into forest soils. Phytopathology 57:1265.

Toussoun, T. A., and Paul E. Nelson.

1968. A pictorial guide to the identification of Fusarium species. 51 p.

Pa. State Univ. Press, College Park.

10
3. CHARCOAL ROOT DISEASE

Macrophomina phaseoli (Maubl.) Ashby

RICHARD S. SMITH, JR.

The charcoal root disease caused by the fungus Macrophomina

phaseoli (Sclerotium bataticola) is one of the most important

diseases in forest nurseries of western North America. It was

originally described as a disease of agricultural crops, but became

important in the 1950's and 1960's, when the forest nursery

program began to expand. The general shift to large, more

efficient forest nurseries in the warmer agricultural areas resulted

in an increase in the charcoal root disease.

Distribution

This disease has worldwide distribution in the temperate and

tropical areas of both hemispheres. It is found in most of the

warmer agricultural areas of the United States. In the South

and Southeast, this fungus, together with other fungi, causes the

black root rot of pine. In the western United States it is a

problem in the warmer lowland agricultural soils, but not in

the cooler, higher elevation forested areas. It is also absent from

the cool coastal areas of the Pacific Coast States.

Hosts and Damage

M. phaseoli attacks more than 300 species of plants including

many agricultural plants, forest tree seedlings, and native weeds.

All conifers are probably susceptible to some degree. Field observations suggest that ponderosa and Jeffrey pines are the least

susceptible while sugar and Monterey pines, Douglas-fir, red and

white fir, and giant sequoia are the most susceptible.

Life History

The fungus invades the root tips, lateral roots, and root crown,

and destroys the cortex, phloem, and cambium. The gradual

destruction of the root system causes the seedlings to become

stunned and chlorotic, and finally to die.

Nursery managers usually feel the impacts of losses in three

ways: (a) seedling mortality in the nursery; (b) increased number of cull (stunted) seedlings; and (c) increased losses of outplanted seedlings due to damaged root systems.

11
The fungus overwinters or lies dormant in the soil in the form

of sclerotia—small, black, spherical resting structures. It does

not grow or develop in the soil in the absence of a host plant.

When the growing root of a susceptible plant comes in contact

with a sclerotium, it is stimulated by the root to germinate. The

fungus grows over the root surface and penetrates between the

epidermal cells into the root cortex. In the cortex the fungus

grows through the root toward the root crown, destroying the

cortex, phloem, and cambium as it proceeds. The fungus continues to develop within these dead tissues, forming masses of

new dormant sclerotia both in the cortex and between the xylem

elements.

These sclerotia can be seen easily in the roots through a hand

lens, and hence are helpful in diagnosis. As other saprophytic

microbes decay the remains of the infected roots, and as soil

movements break up rotten roots, the new sclerotia are liberated

into and distributed within the soil to await the next crop of

seedhngs. These sclerotia may remain dormant for many years

awaiting a suitable host.

Temperature plays a critical part in this disease. The fungus

requires rather warm soils (15°-18° C) before it can become

damaging. In California nurseries, this disease does not occur

until midsummer.

Depending on environmental conditions of the outplanting site,

the fungus may or may not continue to threaten the transplanted

seedlings. In an infested nursery, many seedlings become mildly

infected and never show any symptoms. When outplanted, these

seedlings carry the disease with them. The fungus ceases to

develop and gradually dies in seedlings planted in cool forested

sites. In warm soils, the fungus continues to develop and may

kill the transplant. Infected transplants which are placed under

stress, such as a lack of moisture, are most susceptible.

Control

Most cultural control measures are not effective against this

pathogen. The ability of the sclerotia to survive many years in

the soil and their wide host range limit effectiveness of fallow

periods and cover crops as control measures. In some areas where

soil temperatures are marginal for disease development, seedling

mortality may be reduced by shading, but the general level of

seedling infection is not usually reduced markedly.

The most successful control measures to date are preplant soil

fumigations with broad-spectrum biocides such as methyl bromide

or mixtures of methyl bromide and chloropicrin. Soil fumigation

either in spring or early fall before planting is effective in controlling this disease. Both pre- and post-planting applications of

several fungicides have been tested, but none has done an

adequate job.

12
Selected References

Hodges, Charles S.

1962. Black rot of pine seedlings. Phytopathology 52:210-219.

Seymour, C. P.

1969. Charcoal rot of nursery grown pines in Florida. Phytopathology

59:89-92.

Smith, Richard S., Jr., and Robert V. Bega.

1964. Macrophomina phaseoli in the forest nurseries of California. Plant

Dis. Rep. 48:206.

Smith, R. S., Jr., and R. V. Bega.

1966. Root disease control by fumigation in forest nurseries. Plant Dis.

Rep. 50:245-248.

Smith, Richard S., Jr., and Stanley L. Krugman.

1967. Control of the charcoal root disease of white fir by fall fumigation. Plant Dis. Rep. 51:671-674.

13
4. BLACK ROOT ROT OF PINE

Sclerotium bataticola Taub.

CHARLES S. HODGES*

Black root rot is the most serious root disease of pine seedlings

in southern forest tree nurseries. Mortality of seedlings is not

too great in the nursery as long as adequate water is available,

but survival of outplanted diseased seedlings is low.

Distribution and Hosts

Black root rot has been found in nurseries in most southern

States as far west as Texas, but it occurs most commonly in

southern Georgia and northern Florida. A somewhat similar

disease has been reported from California.

All species of southern pines tested, including slash, loblolly,

longleaf, pond, shortleaf, Virginia, and sand, are highly susceptible to black root rot. Spruce pine appears to be highly resistant.

Symptoms

The earliest symptoms of black root rot are detectable when

the seedlings are about 2 months old. These consist of a slight

enlargement and blackening of the distal portion of the taproot

and larger laterals (fig. 4-1). By midsummer, the lower part of

the taproot and most of the lateral roots may be dead. Numerous

new lateral roots are usually formed above the dead portion of

the taproot, and these may also be killed.

The above symptoms are characteristic of many root diseases.

The distinctive symptom of black root rot is rough, dark enlarged

areas on the taproot and larger laterals. Enlargements are usually

observed about 1 month after the onset of symptoms, and first

appear as small, reddish-brown areas which may remain localized

or eventually may cover the entire taproot. They may reach two

or more times the diameter of the normal taproot, are corky in

texture, very rough, and eventually turn black. Few lateral roots

remain alive on such affected portions of the taproot.

Principal Plant Pathologist, Southeastern Forest Experiment Station,

USDA Forest Service, Research Triangle Park, N.C.

14
F-622496

Figure 4-1.—Roots of a healthy pine seedling (jight) and roots damaged by

black root rot.

The enlargements an the roots are due to the proliferation and

enlargement of the cells of the phelloderm. The phellum of

diseased roots is also many times thicker than on healthy roots,

and this thickening accounts for the roughened appearance of

the root surface.

Aboveground symptoms are not always correlated with root

symptoms. Because the seedlings in the nursery grow under

optimum conditions of moisture and nutrition, it is not unusual

to find seedlings with advanced root symptoms that show no top

symptoms. Chlorosis and stunting may occur in some instances,

however.

15
Life Cycle and Control

The two organisms most commonly associated with black root

rot, Fusarium oxysporum and Sclerotium bataticola (Macrophomina phaseoli), are facultative saprophytes which can survive in

the soil in the absence of their hosts. Both organisms produce

thick-walled resting spores which can persist in the soil for many

years under adverse conditions. Soil fumigation is therefore the

recommended means of control. Methyl bromide or mixtures of

methyl bromide and chloropicrin are the most effective fumigants.

Two nurseries where black root rot became severe the first year

of operation were established on cleared pine sites. This fact,

coupled with the finding of black root rot in a number of pine

plantations in north Florida, indicates that the organisms responsible for the disease are indigenous to forest areas. Thus nurseries should not be established on newly cleared forest land.

With one exception, all nurseries where black root rot has been

found are located on light, sandy soil.

Selected References

Hodges, Charles S.

1962. Black root rot of pine seedlings. Phytopathology 52:210-219.

Rowan, Samuel J.

1960. The susceptibility of twenty-three tree species to black root rot.

Plant Dis. Rep. 44:646-647.

16
5. PHYTOPHTHORA ROOT ROT

Phytophthora cinnamomi Rands

E. G. KUHLMAN AND RICHARD S. SMITH, JR.*

In the past, Phytophthora root rot has caused losses in many

species of coniferous and hardwood seedlings throughout the

United States. In recent years, however, soil fumigation prior to

planting has reduced the loss to this disease in most areas, except

where seedlings are held in nursery beds for several years. The

concern over the possible spread of this disease to new areas is

apparent from some States' quarantine regulations forbidding

intercounty shipment of infected stock.

Distribution and Hosts

Phytophthora cinnamomi has been isolated from soil and from

forest and ornamental plants throughout the United States. It

is most common in forest and agricultural lands in the southeastern United States and in the warmer agricultural lands and

forest nurseries in western North America. Recent surveys indicate, however, that the fungus is present in only a few forest

nurseries. It probably is absent from our western forests.

^ Over 100 species of plants have been reported as hosts for P.

cinnamomi In forest nurseries, major losses have been experienced in red pine, Fraser fir, black walnut, and yellow-poplar.

The wide distribution and large host range make this fungus a

potential threat in many nurseries.

Symptoms

Symptoms in nurseries often are not evident until seedlings

start dying, since infection of the host plant commonly occurs

through the feeder roots. Infected plants may remain symptomless until the tops wilt. They then rapidly turn a grey-greenbrown and die. In some nursery beds this disease appears as

enlarging circles of chlorotic and stunted seedlings, with the most

severely affected seedlings at the center. In most hosts the disease

is characterized by a red-brown discoloration of the cambial

region, particularly at the root collar and sometimes extending

several inches up the stem. In black walnut and chestnut, this

discoloration is a blue-black "inky" color. The characteristic redbrown color is also present in the cambium and wood or roots, so

that diseased seedlings can be identified prior to the appearance

of top symptoms. In some hosts only root tips are killed, which

results in chlorosis and stunting.

»Respectively, Principal Plant Pathologist, Southeastern Forest Experiment Station, USDA Forest Service, Research Triangle Park, N.C. and

Principal Plant Pathologist, Pacific Southwest Forest and Range Experiment

Station, USDA Forest Service, Berkeley, Calif.

17
Life Cycle

Chlamydospores or oospores formed in host tissue enable the

fungus to survive for years in the soil. When fresh host material

becomes available, these spores germinate and produce motile

zoospores or infective mycelium. Most commonly, zoospores swim

a few inches or are carried in ground water to the root tip.

Infection occurs in the region of elongation or through wounds.

Chlamydospores, oospores, and more zoospores are formed soon

after infection. The former provides means of long-term survival

whereas zoospores provide rapid spread along the root or to

adjacent roots. Depending on the host species, the fungus may

spread up through the roots to the root crown, killing the

cambium and inner bark as it grows, or it may be limited to the

root tips.

High soil moisture levels are necessary for zoospore formation

and release. Soil temperatures above 15° C are necessary for

infection and colonization of the host. These conditions are common in most nurseries.

Control

Fumigation of the soil prior to planting is the most certain

control for Phytophthora root rot. If stock is only in the nursery

for 1 year, soil fumigation with combinations of methyl bromide

and chloropicrin will prevent this disease. When stock is held

in a nursery for several years, extreme care should be taken to

prevent introduction of this pathogen. Any stock or ornamentals

brought into the nursery should be thoroughly checked and certified disease free. If the pathogen is present in a nursery, immediate fumigation of the infested area and a 2- to 3-foot surrounding buffer zone is necessary. Any effort to save apparently

healthy stock in the infested area will result in the spread of the

fungus to other areas of the nursery, as well as to outplanting

areas. Particular care should be taken to treat border areas that

may have been contaminated from soil carried on equipment. All

equipment should be thoroughly cleaned before proceeding from

the infested area. Soil drenches have not been effective in control

of the disease, probably because chlamydospores and oospores

are not affected by the drenches.

Selected References

Hendrix, Floyd F., Jr., and W. A. Campbell.

1968. Pythiaceous fungi isolated from Southern forest nursery soils and

their pathogenicity to pine seedlings. For. Sei. 14:292-297.

Hodges, Charles S., Jr.

1962. Diseases in Southeastern forest nurseries and their control. U.S.

Dep. Agrie. For. Serv. Southeast For. Exp. Stn. Stn. Pap. 142, 16 p.

Kuhlman, E. George, and Floyd F. Hendrix, Jr.

1963. Phytophthora root rot of Fraser fir. Plant Dis. Rep. 47:552-553.

Thorn, William A., and George A. Zentmyer.

1954. Hosts of Phytophthora cinnamomi Rands. Plant Dis. Rep. 38:47-

52.

18
6. PHYTOPHTHORA ROOT ROT OF BLACK

WALNUT SEEDLINGS

Phytophthora sp,

RALPH J. GREEN, JR.*

Black walnut seedlings are highly susceptible to a root rot

disease in the seedbed caused by fungi of the genus Phytophthora.

Infection may occur at any time after seeds begin to germinate.

Early infection reduces emergence and usually kills young seedImgs. Older seedlings may survive, but the disease continues to

develop in overwinter storage and may cause serious losses in

the spring during handling and shipping.

Distribution

Phytophthora root rot occurs wherever black walnut seedlings

are grown under nursery conditions. It is apparently limited to

nursery seedbeds, however, and is not a recognized problem in

natural regeneration.

Hosts and Damage

Although black walnut is highly susceptible to Phytophthora

root rot, the disease is not confined to this host species. It also

occurs on seedlings of the oaks, American and oriental chestnuts,

and numerous other hardwood and coniferous species.

Infection may occur at any time after the seed splits prior to

germmation. If the germinating seed is invaded, the seedling is

killed before it emerges above ground. Young seedlings that are

infected usually collapse suddenly, blacken, and die in a few days.

Older seedlings usually wilt and yellow before the plants are

killed. These plants also turn black and the leaves hang on.

The root system of diseased plants shows extensive brown,

water-soaked lesions (fig. 6-1) that begin at the root collar or at

the point of attachment of a lateral root. The root rot progresses

rapidly and may involve the lower stem as well as the root system.

Infected seedlings usually occur in spots or patches in the seedbed, which suggests that the fungus spreads from plant to plant

after infection begins. Root rot is always more severe in low

areas of the plant beds where drainage is poor.

^Professor of Plant Pathology, Plant Pathology Department, Purdue

University, W. Lafayette, Ind.

19
F-622497

Figure 6-1.—Symptoms of root rot of black walnut seedlings caused by

Phytophthora citricola.

20
When walnut seedlings are lifted in the fall, many plants have

dead or discolored lateral or fibrous roots and often show restricted discolored areas in the taproot cortex. After winter storage, these infected seedlings may develop root rot symptoms very

rapidly during handling and shipping.

Symptoms on the oaks and chestnuts are similar to those on

black walnut, except that infected seedlings turn brown or tan

rather than black. Also, these species are apparently much less

susceptible to this root rot than are walnut seedlings.

There is little evidence that this disease is a continuing problem

after disease-free walnut seedlings are outplanted. However,

seedling stock that is not carefully inspected and graded or that

is improperly handled may be seriously reduced in vigor, and

mortality may be high after transplanting.

Life History

Phytophthora root rot of black walnut seedlings and other tree

species may be caused by any one of at least three different

species of this fungus. These include Phytophthora cinnamomiy

P. cactorum and, more recently, P. citricola. The symptoms produced and the general disease cycle for all three fungi are similar.

The causal fungi are soilborne, and persist indefinitely in soil

on plant debris and other organic matter. They are considered

weak parasites and are usually limited to very young seedlings

or plants that have been damaged or weakened. These fungi are

also favored by high soil moisture for active migration of their

motile spores.

Experimental studies have shown that one or more of these

fungi is present in virtually all nursery soils. Conditions favorable for infection include heavy soils, high water tables, or heavy

rains which saturate the soil for periods of several days. The

fibrous roots of seedlings are damaged or killed by lack of oxygen

in saturated soils, and the root rot fungi rapidly colonize the

injured root tissues. These fungi then grow into the sound roots

and lower stems, where they kill young seedlings rapidly, or

produce root infections in older seedlings. The disease is most

common under cool temperatures, but soil temperature and other

environmental factors are not so important in the infection

process as high soil moisture.

Control

Attempts have been made to control this disease by soil fumigation and by various chemical treatments of soil and seed.

Although root rot incidence may be somewhat reduced, there is

no completely effective chemical control.

Walnut root rot can be controlled to some degree by avoiding

areas of poor drainage. The nursery area should be tiled and well

drained for effective water management.

21
Seedlings should be carefully inspected and graded when lifted

in the fall for winter storage. Bulk storage in cold rooms should

be avoided because these fungi grow rapidly at rather low temperatures. After overwinter storage, walnut seedlings should not

be held at temperatures above 4° C for periods longer than

necessary to process and ship the seedlings.

Selected References

Crandall, Bowen S., G. F. Gravatt, and Margaret Milburn Ryan.

1945. Root disease of Castanea species and some coniferous and broadleaf nursery stocks caused by Phytophthora cinnamomi. Phytopathology 35:162-180.

Crandall, Bowen S., and Carl Hartley.

1938. Phytophthora cactorum associated with seedling diseases in forest

nurseries. Phytopathology 28:358-380.

Green, R. J., Jr., and R. G. Pratt.

1970. Root rot of black walnut seedlings caused by Phytophthora

citricola. Plant Dis. Rep. 54:583-585.

Waterhouse, Grace M.

1963. Key to the species of Phytophthora de Bary. Commonwealth

Mycol. Inst., Mycol. Pap 92, 22 p.

22
7. CYLINDROCLADIUM ROOT ROT

Cylindrocladium scoparium Morgan,

C. floridanum Sobers & Seymour

CHARLES E. CORDELL AND DARROLL D. SKILLING*

Cylindrocladium root rot causes severe losses to both hardwood

and conifer seedlings in forest tree nurseries. The problem has

increased in the past few years due to internursery movement

of infected stock. The damage was formerly attributed only to the

fungus Cylindrocladium scoparium. However, a closely related species, C. floridanum, is also a part of this disease complex.

F-522499

Figure 7-1.—Cylindrocladium (C. floridanum) root rot damage (approximately

70 percent) to 1-0 yellow-poplar seedlings in a North Carolina forest tree

nursery.

*Respectively, Supervisory Plant Pathologist, Forest Pest Management,

Division of Environmental Protection and Improvement, Southeastern Area,

USDA Forest Service, Asheville, N.C., and Principal Plant Pathologist,

North Central Forest Experiment Station, USDA Forest Service, St. Paul,

Minn.

23
Distribution

The disease is well distributed in nurseries throughout the

eastern, southeastern, and Lake States regions. Although the

disease is not reported in western nurseries, many western tree

species are known to be susceptible to the fungus. The disease has

also been reported from one forest nursery in Quebec.

Hosts and Damage

C. scoparium and C. floridanum have a wide host range. The

disease caused by these fungi has been reported on plants in 66

genera of 31 families in many areas of the world. Susceptible

conifer tree species include Austrian, jack. Mugo, red, Scotch, and

white pines ; black, blue, Norway, and white spruces ; balsam and

Fraser firs; and Douglas-fir. The most serious losses have occurred on red and white pines and black and white spruces.

Yellow-poplar and black walnut are the most commonly infected

and the most severely damaged hardwood species (fig. 7-1).

Cylindrocladmm spp. cause several kinds of damage, including

root rot, damping-off, needle and leaf blight, and stem cankers.

Root Rot

This is the most common symptom observed. Infected conifer

roots become necrotic and discolored, and the cortex is easily

removed in advanced stages of rot. The roots die from the tips

back toward the root crown. New lateral roots may develop prolifically above the infected area, usually just below the root

crown. Aboveground symptoms of conifer root rot are yellowing

of needles as the seedlings enter periods of moisture stress. The

foliage gradually becomes reddish brown as moisture loss progresses (fig. 7-2).

Roots of infected hardwood seedlings, such as yellow-poplar,

develop a pronounced blackened condition and longitudinal cracking of the root cortex in contrast to the smooth white roots of

healthy yellow-poplar seedlings (fig. 7-3).

Damping-Off

This disease occurs on newly emerging seedlings. Infected conifer stems above and below the ground become shrunken, dry, and

turn brick red. In advanced stages, the seedling top can be lifted

from the soil surface because the root system has been destroyed.

Because other damping-off fungi cause similar symptoms, a laboratory diagnosis is necessary for positive identification of the

pathogen.

Needle and Leaf Blight

On red pine, infection occurs at the base of needle fascicles

during moist weather ; new needles are the most susceptible. The

24
Ifffttti?hff )ít(M

'*-I»

1 "^ ■■ "«^

F-621562

Figure 7-2.—Cylindrocladium root rot

on 2-1 red pine in a Wisconsin

nursery.

F-B22501

Figure 7-3.—C. floridanum root rot

symptoms on a 1-0 yellow-poplar

seedling. Note the blackened and

longitudinally cracked cortex of the

infected root.

infected needles turn yellow and develop brown lesions that

enlarge, become sunken, and darken. The needles finally turn

brick red and fall off. On white pine, infected needles yellow from

their tips toward the base, and the brick red color is not as

conspicuous. On black, Norway, and white spruces, the foliage

turns yellow and the new growth may wilt.

Infected hardwood foliage shows necrotic leaf spots. Many hardwood seedlings that have severe leaf blight symptoms die.

Stem Cankers

Stem cankers have been most commonly observed on white

pine seedlings, generally following the needle blight stage. They

may occur randomly along the seedling stem, but usually originate at the base of needle fascicles. Brownish, sunken areas

gradually enlarge, may girdle the stem, and kill the seedling.

Life History

The causal fungus overwinters in the infected roots or needles

as thick-walled resting bodies called microsclerotia. Under favorable environmental conditions the microsclerotia germinate, and

germ tubes invade the hypocotyls of germinating seeds or the

roots of seedlings or transplants. Airborne conidia are produced

25
during the growing season on infected foliage. During hot, humid

weather the conidia germinate and their germ tubes enter the

needle stomates. This results in the needle blight stage. Resting

bodies produced within and/or on infected needles, roots, and

soil organic matter contribute to soil infestation when dead

needles fall to the soil surface and when invaded roots die.

Control

The intensity of Cylindrocladium root rot can be significantly

reduced by soil fumigation to a depth of at least 12 inches before

seedmg or transplanting. Use a soil fumigant containing 67 percent methyl bromide and 33 percent chloropicrin at a rate of at

least 300 pounds of the product per acre. Cover the: fumigated

beds immediately with plastic tarping. DMTT (Mylone) at 200

to 250 pounds per acre has been used in several nurseries ; however, control has not been quite as good as with methyl bromide.

Because no tarping is used, a fumigant such as DMTT is effective only to the depth that it is mixed with the soil.

Bordeaux mixture has been used as a foliage spray to prevent

needle blight of seedlings.

Selected References

Cordell, C. E., A. S. Juttner, and W. J. Stambaugh.

1971. Cylindrocladium floridanum causes severe mortality of seedling

yellow-poplar in a North Carolina nursery. Plant Dis. Rep. 55:700-

Cox, R. S.

1954. Cylindrocladium scoparium on conifer seedlings. Del A^ric Exn

Stn. Tech. Bull. 301, 40 p.

Filer, T. H., Jr.

1970. Virulence of three Cylindrocladium species to yellow-poplar seedlings. Plant Dis. Rep. 54:320-322.

Hodges, Charles S., Jr.

1962. Diseases in southeastern forest nurseries and their control U S

Dep. Agrie. For. Serv., Southeast. For. Exp. Stn., Stn. Pap. 142!

16 p. 1- »

Sobers, E. K., and C. P. Seymour.

1967. Cylindrocladium floridanum sp. n. associated with decline of peach

trees m Florida. Phytopathology 57:389-393.

Thies, Walter G., and Robert F. Patton.

1970. The biology of Cylindrocladium scoparium in Wisconsin forest

tree nurseries. Phytopathology 60:1662-1668.

26
8. CROWN GALL

Agrobacterium tumefaciens (E. F. Smith & Towns.)

Conn.

GLENN W. PETERSON AND LARRY W. MOORE*

Crown gall, caused by the bacterium Agrobacterium tumefaciens, is an economically important disease in commercial nurseries, especially those growing numerous hardwood species. Although this disease has so far been of minor importance in

forest tree nurseries, it warrants attention because of its potential

for economic loss and the difficulty in controlling the disease.

Distribution and Hosts

Crown gall disease is widespread, occurring on this and other

continents. A wide range of trees are susceptible, including both

conifer and broadleaf species. Natural infection of species of

F-522B03

Figure 8-1.—Crown gall that developed on eastern redcedar root following

root pruning.

*Respectively, Principal Plant Pathologist, Rocky Mountain Forest and

Range Experiment Station, USDA Forest Service, Lincoln, Nebr., and

Assistant Professor of Plant Pathology, Department of Botany and Plant

Pathology, Oregon State University, Corvallis, Ore.

27
Juniperus, Cupressus, and Libocedrtis has been reported. There

seems to be no report of infection of pines in nurseries. Several

Abies species have been infected following artificial inoculation.

Two strains of A. tumefaciens have been isolated from naturally galled plants. The strains are most conveniently classified by

whether they produce a ketoglycoside from lactose, but Keane et

al. (1970) have listed a number of other biochemical properties

which the strains do not have in common.

Recent work in Australia and Oregon has shown that the

auxotrophic ketoglycoside-negative strain of the bacterium has

been responsible for most infections occurring in those two geographic areas. The ketoglycoside-negative strain has been isolated

from galled Prunus and Rubus species, willow, roses, apples,

Norway maple, and soil from nurseries. Both ketoglycoside-negative and ketoglycoside-positive strains have been simultaneously

isolated from naturally occurring galls on birch and mountain ash.

This repeated isolation of the ketoglycoside-negative strain is

curious inasmuch as the prototrophic ketoglycoside-positive strain

has been most commonly used in research and is the predominant strain maintained in most culture collections. It may be because isolation procedures were used which selected for the ketoglycoside-positive strain.

Description of Damage

Infected plants usually develop galls (tumorlike swellings) on

roots and stems below ground (fig. 8-1). On some species, such

as cottonwood and willow, galls are also common above ground on

stems. The galls may be a few to several times the diameter of

the root or stem tissue on which they develop.

Nursery losses are incurred primarily because infected stock

must be culled. Inspection agencies do not permit the distribution

of stock containing galls. Such losses vary from 10 to 100 percent

of a given nursery crop.

F-522604

Figure 8-2.—Natural infection of crown gall on nursery-grown mazzard cherry

seedlings. Note common point of formation of galls.

28
The adverse influence of infection on some species grown for

fruit or nuts has been established; however, little or no such

information is available for susceptible forest trees. Early infection of young ornamental flowering cherry seedlings at critical

points such as the crown can greatly reduce root numbers and

stunt the tree (fig. 8-2).

Life History

Crown gall bacteria are apparently soilborne. They have been

reported to persist in the soil from 1 to 2 years in the absence

oí a host. Because these bacteria enter plants through manmade

or naturally occurring wounds, the common nursery practice of

rootpruning encourages infection. Eastern redcedar in a central

Nebraska nursery had galls develop at the point where roots

had been pruned. Galls are commonly observed on the pruned end

of taproots in hardwood nursery stock.

Natural wounds caused by chewing insects (such as grubs and

wire worms), nematodes, and lateral root initiation have also been

implicated as aiding infection by crown gall bacteria.

The crown gall bacteria are easily spread and dispersed by

contaminated irrigation or runoff water that has passed through

orchards containing galled trees. This source should be considered

when watering stratified seed that is being planted in the spring.

^ Shipment of diseased planting stock obviously contributes to

dispersal of the crown gall bacteria to other geographic areas.

Although galled trees are usually detected easily, latent infections

have been shown to occur in several Prunus species and in apples.

Latent infections are diflficult, if not impossible, to detect by visual

inspection. The problem of recognizing latent infections in interstate shipment of nursery stock is compounded because of

differing environmental conditions (especially cool vs. warm soil

and air temperatures) from one geographic area to another which

affect symptom expression.

Control

Fumigants have reportedly killed crown gall bacteria in host

tissue in the soil when used at rates about twice that needed for

nematode control. Control by fumigants has not been consistent,

however. The disease was not controlled in beds in a Plains nursery which were fumigated with methyl bromide to control

nematodes. In some cases, soil fumigation has resulted in increased infection.

Because the bacterium can reportedly overwinter in the soil

m the absence of a susceptible host, beds containing stock with

crown gall should be replanted with an immune or highly

resistant species. Populations of crown gall bacteria probably

are reduced by using nonsusceptible cover crops (grain crops)

in rotations. However, the possibility exists that the bacteria

could survive as saprophytes in the rhizosphere and rhizoplane of

29
nonsusceptible hosts. Several growers of ornamental Prunus

species observed a relatively high incidence of crown gall when

they replanted as long as 4 years later to fields in which a

severe crown gall problem had previously occurred.

The effectiveness of preplanting treatments in preventing

crown gall infection has been sporadic, perhaps due to the possibility of latent infections and subsequent inoculation and mfection as a result of growth cracks and wounds sustained at a

latpr neriod

Because crown gall disease is highly contagious, good sanitary

practices and pathogen-free planting stock are a must to reduce

the incidence of disease. Tools and equipment, grading tables,

and seed beds (if possible) should be surface sterilized. Nursery

stock should be carefully handled, planted, cultivated, harvested,

and stored to minimize wounding of susceptible tissue.

Irrigation water from deep wells should be free of crown gall

bacteria, but water from irrigation lines or ditches that have

passed through fields or plots with galled trees should be avoided.

Similarly, any cultural practice that places healthy trees with

galled trees in standing water is very risky.

Selected References

Bernaerts, M. J., and J. DeLey.

1963. A biochemical test for crown gaU bacteria. Nature 197:406-407.

Deep, Ira W., Ray A. McNeilan, and Iain C. MacSwan.

1968. Soil fumigants tested for control of grown gall. Plant Dis. Rep.

52:102-105.

Deep, Ira W., and Roy A. Young. , . .

1965. The role of preplanting treatments with chemicals in increasing

the incidence of crown gall. Phytopathology 55:212-216.

Dickey, Robert S. , ^ ^ r ^ •

1962. Efficacy of five fumigants for the control of Agrobacterium

turnefadens at various depths in the soil. Plant Dis. Rep. 46:73-76.

Keane, P. J., A. Kerr, and P. B. New.

1970. Crown gall of stone fruit. II. Identification and nomenclature

of Agrobacterium isolates. Aust. J. Biol. Sei. 23:585-595.

1969. Crown gall of stone fruit. I. Isolation of Agrohacterium tumefaciens and related species. Aust. J. Biol. Sei. 22:111-116.

Munnecke, Donald E., and John Ferguson. - . ^. -

1960. Effect of soil fungicides upon soil-borne plant pathogenic bacteria

and soil nitrogen. Plant Dis. Rep. 44:552-555.

Riker, A. J., and E. M. Hildebrand. „ ^ ,

1934. Seasonal development of hairy root, crown gall, and wound overgrowth on apple trees in the nursery. J. Agrie. Res. 48:887-912.

Schroth, M. M., A. R. Weinhold, A. H. McCain, D. C. Hildebrand, and N.

1971. Biology and control of Agrohacterium tumefaciens, Hilgardia 40:

537-552.

Smith, Clayton 0. n j 4.

1939. Susceptibility of species of Cuppressaceae to crown gall as determined by artificial inoculations. J. Agrie. Res. 59:919-925.

Smith, Clayton 0.

1942. Crown gall on species of Taxaeeae, Taxodiaeeae, and Pmaeeae, as

determined by artificial inoculation. Phytopathology 32:1005-1009.

30
9. NEMATODES

JOHN L. RUEHLE*

Nursery seedlings are particularly vulnerable to soilborne

nematode diseases. Continuous cultivation of the same or similar

plant species within restricted areas is conducive to the rapid

buildup of parasitic nematode populations, and if not controlled,

nematode diseases can result in significant losses. Such losses

reduce the economic value of nursery crops of both conifers and

hardwoods. Additional sales may also be lost because regulations

restrict shipments of nematode-infected plants.

Distribution and Hosts

Plant-parasitic nematodes are found in forest nurseries

throughout North America. Root-knot nematodes are not usually

important parasites in areas with a cool climate. Conversely, certain species of lesion nematodes, serious in cooler regions, are

rarely found in the southern United States.

Some parasitic nematodes have a wide host range and feed on

many forest tree species; others are quite restrictive in their

feeding habits. The pine cystoid nematode feeds only on certain

conifers, and several species of root-knot nematodes infect certain hardwoods but not conifers. The nematodes most damaging

to forest nursery crops are root-knot nematodes, Meloidogyne

spp. ; pine cystoid nematodes, Meloidodera spp. ; lance nematodes

Hoplolaimus spp.; lesion nematodes, Pratylenchus spp.; stunt

nematodes, Tylenchorhynchus spp.; stubby-root nematodes, Trichodorus spp. ; and dagger nematodes, Xiphinema spp.

Description of Damage

The reaction of plants to attack by plant-parasitic nematodes

varies considerably with the host parasite combination. Nematode diseases ordinarily cannot be diagnosed by symptoms alone

because other parasitic organisms or certain environmental factors can cause similar damage to roots. The aboveground symptoms of nematode diseases of forest nursery seedlings are

generally similar and are usually the same as symptoms on any

plant which has been deprived of a properly functioning root

system. Diseased seedlings are stunted, lack vigor, and their

*Principal Plant Pathologist, Forestry Sciences Laboratory, Southeastern

Forest Experiment Station, USDA Forest Service, Athens, Ga.

31
leaves are reduced in size and number and become chlorotic;

they often show symptoms of mineral deficiency, even when adequate levels of soil fertility are maintained. Affected plants

usually cannot withstand long periods of low soil moisture.

Nurseries with nematode disease usually have patches of

stunted seedlings, characterized by low vigor, having small, distorted root systems (fig. 9-1). These patches enlarge and coalesce

as nematode populations increase and spread through the nursery

beds.

The belowground symptoms appear as root. impairment or

distortion. Root-knot nematodes may cause galling of roots and

in some cases a proliferation of roots near the galls. Dagger

nematodes on certain hosts cause apical root-galling and distortion of lateral roots. Pine cystoid nematodes cause little, if any,

galling and are apparent only when the swollen females erupt

through the root epidermis. In contrast, lesion nematodes cause

a necrosis of'the cortex resulting in a general decay of the feeder

roots. The lance nematode, an endoparasite particularly damaging to pine seedlings, also migrates through the cortical tissue

and causes extensive internal cell destruction. This type of damage by internal parasites often allows fungus pathogens to enter

and accentuate the general decay of the root system.

Nematodes that do not enter the roots generally have less

obvious effects on forest tree seedlings. Stunt, stubby-root and

dagger nematodes cause discoloration and surface lesions, but

more often the effect of their feeding is a general stunting of

lateral roots without noticeable decay (fig. 9-2).

F-B22505

Figure 9-1.—Root lesion nematode damage to ponderosa pine seedlings, and

control by methyl bromide fumigation.

32
Figure 9-2.—Comparison of healthy root system of sand pine seedling (left),

and poorly developed root system caused by parasitic nematodes (right).

Nematodes often interact with fungi to produce disease complexes of tree seedlings. Certain rhizosphere fungi are destructive only when they occur in combination with nematodes parasitizing feeder roots. The damage caused by certain fungus

pathogens capable of infecting feeder roots is often increased by

joint action with plant-parasitic nematodes. Certain nematodes

also prevent normal synthesis of mycorrhizal development on

feeder roots by either directly attacking the fungus symbiont or

damaging root tissue before mycorrhizal synthesis.

Many soilborne viruses are transmitted by nematodes. Xiphinema spp. and Longidorus spp. are known vectors of certain

viruses that infect tree seedlings.

Control

Since nematodes cannot be eradicated in nursery soils even

with broadcast fumigation, nematode control in nurseries is

based on the concept of lowering nematode populations to levels

below which yield is not affected.

Soil fumigation with methyl bromide at rates of 300-500

pounds per acre gives excellent control of nematodes in forest

nurseries. Methyl bromide produces additional benefits by controlling insects, bacteria, fungi, and weeds and by increasing

the availability of certain soil nutrients. Dowfume MC-33 and

Brozone, general purpose fumigants containing methyl bromide.

33
also give excellent nematode control. When nematodes are a

problem in small areas in a nursery, spot injection with less

volatile nematicides, such as DD mixtures, will give satisfactory

control.

Recently, bare roots of infected hardwoods have been dipped

in chemicals to control root-knot nematodes. Organophosphate

nematicides, such as Zenophos, have satisfactorily controlled

nematodes on dogwood. Hot water treatment (52° C for 2 minutes to bare roots) gives good control of lesion nematodes infecting eastern redcedar.

Crop rotation is recommended for the control of certain nematodes. For example, if root-knot nematode is the problem, rotation with fescue (a nonhost) is recommended over soybean (a

favorable host). Nonhost seedlings may be rotated with susceptible seedlings if space is limited and production must be maintained.

Selected References

Christie, Jesse R.

1959. Plant nematodes—their bionomics and control. 256 p. Fla. Agrie.

Exp. Stn., Gainesville.

Peterson, Glenn W.

1962. Root lesion nematode infestation and control in a plains forest

tree nursery. U.S. Dep. Agrie. For. Serv. Rocky Mt. For. and

Range Exp. Stn. Res. Note 75, 2 p. Fort Collins, Colo.

Ruehle, John L., and J. N. Sasser.

1962. The role of plant-parasitic nematodes in stunting of pines in

southern plantations. Phytopathology 52:56-68.

Ruehle, John L., Jack T. May, and S. J. Rowan.

1966. Nursery fumigation trial with Vorlex. Tree Planters* Notes 76,

p. 4-7.

Sutherland, Jack R., and R. E. Adams.

1966. Population fluctuations of nematodes associated with red pine

seedlings following chemical treatment of the soil. Nematologica

12:122-128.

34
10. MYCORRHIZAE OF FOREST NURSERY

SEEDLINGS

DONALD H. MARX*

All species of trees growing under normal soil conditions in

the forest are dual organisms—part plant and part root-inhabiting fungi. These highly specialized, root-inhabiting fungi are not

pathogenic and, thus, do not cause root disease. They are beneficial to their tree hosts. They infect, symbiotically, the cortical

tissues of young feeder roots of trees and form mycorrhizae.

The word mycorrhizae means "fungus-roots." Mycorrhizae are

the normal feeder roots of all forest trees, as they are for the

vast majority of other plant species, including agronomic, horticultural, ornamental, and turf crops.

Classes of Mycorrhizae

The three classes of mycorrhizae are identified by the arrangement of the hyphae of the fungi within root cortical tissues.

Ectomycorrhizae.—This class occurs normally on roots of

pine, spruce, fir, beech, birch, eucalyptus, alder, oaks, hickories,

and certain other trees. Ectomycorrhizae usually can be distinguished from nonmycorrhizal roots because of their swollen

appearance and in the case of Pinus, a forking habit (fig. 10-1).

F-522B07

Figure 10-1.—Three different ectomycorrhizae of Pinus taeda seedlings. Note

branching habits and color differences. Each type was formed by a different species of ectomycorrhizal fungus. Unbranched ectomycorrhizal roots on

right segment are very similar to nonmycorrhizal roots.

*Principal Plant Patholog-ist, Forestry Sciences Laboratory, Southeastern

Forest Experiment Station, USDA Forest Service, Athens, Ga.

35
Not all forked feeder roots of pine are ectomycorrhizal, however, since feeder roots of many species can be stimulated to

fork by many factors other than ectomycorrhizal fungus infection. The fungi which form ectomycorrhizae are among those

that mainly produce mushrooms and pufïballs periodically

throughout the year. Over 2,400 species of ectomycorrhizal fungi

are found in North America alone.

Under microscopic examination, hyphae of ectomycorrhizal

fungi can be observed growing internally only around the cortical cells of the root forming the Hartig net, thus the prefix

ecto. The Hartig net hyphal arrangement appears to replace the

middle lamella, which is normally composed of pectins, and

cements the cortical cells. Hyphae of the fungal symbionts that

are present around the outside of the feeder root usually are

tightly interwoven, and are designated the fungus mantle. The

mantle of ectomycorrhizae may be only one or two hyphal

diameters in thickness, or several dozen hyphal diameters. Ectomycorrhizae may be white, brown, yellow, black, blue, or blends

of these colors. The color of ectomycorrhizae is apparently determined by the hyphal color of the specific mycorrhizal fungus

encompassing the root.

Endomycorrhizae,—This class occurs normally on roots of

maple, sycamore, ash, sweetgum, walnut, cypress, cedar, some

poplar, and certain other hardwoods. Endomycorrhizae cannot

be readily distinguished from nonmycorrhizal roots without the

aid of a microscope. Endomycorrhizal infection does not normally cause color or physical changes in the root. The fungi which

form endomycorrhizae are different from those of ectomycorrhizae and lack conspicuous aboveground fruiting bodies.

Most endomycorrhizal fungi of forest trees apparently belong to the genus Endogone, Dozens of species of endomycorrhizal fungi have been identified; undoubtedly many more exist.

Under microscopic examination, hyphae of the endomycorrhizal

fungi are observed growing into (thus, prefix endo) the cortical

cells of the roots forming arbuscules. Arbuscules are specialized

absorbing hyphae or haustoria of these fungal symbionts. Large

vesicles, which are swollen hyphae, may also be seen in endomycorrhizal roots. If both vesicles and arbuscules are present, the

endomycorrhizae are called "vesicular-arbuscular'' (VA) niycorrhizae. The Hartig net and fungus mantle which characterize

ectomycorrhizae are not present in endomycorrhizae. Endomycorrhizal fungi will frequently produce large, conspicuous, pearlcolored spores (either zygospores or chlamydospores) on hyphae

attached to endomycorrhizal roots (fig. 10-2). The presence of

these spores on root surfaces, which are detectable under low

magnification, is a reasonably reliable indication of endomycorrhizal fungus infection of the root.

Ectendomycorrhizae.—This class is apparently an intermediate between the other two classes. These fungi grow into the

36
F-B22B08

Figure 10-2.—Spores of an endomycorrhizal fungus formed on endomycorrhizae of sweetgum. The diameter of these spores is approximately 0.004

inch. Note hyphae of the fungus radiating from the root surface.

cortical cells of the root with an appearance quite different from

the arbuscular formation of endomycorrhizae. These fungi grow

around the cortical cells in a Hartig net arrangement. They may

or may not develop a fungus mantle over the feeder root surfaces.

Ectendomycorrhizae have been observed on roots of certain

species of tree seedlings in nurseries, especially in the Pacific

Northwest and Northeast, but their significance is unknown. The

species of fungi which form this class of mycorrhizae are also

unknown at present.

Benefits of Mycorrhizae to Trees

A considerable amount of research has been done on the

benefits of ectomycorrhizae to trees (ñg. 10-3). Only limited

research has been done on the importance of endomycorrhizae to

plant growth, and research on benefits of ectendomycorrhizae to

trees is nearly nil. The following list of benefits of mycorrhizae

to tree growth is based almost exclusively on work with ectomycorrhizae; however certain of these benefits are thought to be

derived from endomycorrhizae as well.

1. Tremendous physical increase in absorbing surface of root

system; includes both mycorrhizae and hyphae growing from

mycorrhizae into soil.

2. More selective ion absorption and accumulation, especially

phosphorus.

3. Solubilization of normally nonsoluble minerals and their

constituents.

37
F-622509

Figure 10-3.—Growth dififerences between ectomycorrhizal and nonmycorrhizal

Virginia pine seedlings. The 4-month-old seedlings were well fertilized

and watered.

4. Increased longevity of feeder root function; mycorrhizal

roots persist longer on root systems than do nonmycorrhizal

roots.

5. Resistant to feeder root infections caused by pathogens,

such as Phytophthora and Pythium spp., present in many forest

and nursery soils. This benefit apparently applies only to ectomycorrhizae.

6. Increased tree tolerance to soil toxins (inorganic and

organic), extremes of soil acidity, and high soil temperatures.

General Considerations

1. A given tree species may enter into mycorrhizal association with one or many different species of mycorrhizal fungi at

a given time.

2. Some species of mycorrhizal fungi have very broad treehost ranges, whereas, others have very narrow host ranges.

3. Some species of mycorrhizal fungi are more beneficial to

tree survival and growth than others.

4. Some trees, in particular Pinus, have an obligate need for

mycorrhizae in order to survive. This may not be true for all

tree species, even though mycorrhizae occur normally on their

roots.

5. Certain mycorrhizal fungi are more ecologically adapted

to certain forest sites than are other fungi; trees with adapted

fungal symbionts on roots grow better than trees with nonadapted fungal symbionts growing on the same site.

38
6. Spores of ectomycorrhizal fungi are produced above

ground and are readily wind disseminated. Spores of endomycorrhizal fungi are produced underground and are not wind

disseminated.

7. Many species of ectomycorrhizal fungi can be grown in

pure culture or artificial media. Endomycorrhizal fungi cannot

be grown in pure culture in the absence of their plant hosts.

8. Mycorrhizal fungi rarely exist in an active physiological

state in soil in the absence of their hosts. However, they may remain in a dormant condition, as spores or resistant hyphae, in

soil for many years without a tree host.

9. Not all species of mushrooms or puffballs are ectomycorrhizal. Many are saprophytic in nature and decompose organic

matter.

Practical Considerations

Soil fumigation is becoming more and more a routine practice

in tree nurseries to control weeds, nematodes, and fungal root

pathogens. Most soil fumigants also eradicate the beneficial

mycorrhizal fungi. The eradication of ectomycorrhizal fungi

from nursery soil is usually not a problem since these fungi

produce wind-disseminated spores periodically throughout the

year which recolonize the soil. One of the major ectomycorrhizal

fungi to colonize fumigated nursery soils, especially in the south,

is Thelephora terrestris (fig. 10-4).

r-B22510

Figure 10-4.—Basidiocarp of Thelephora terrestris formed around stem of a

pine seedling. This fungus is ectomycorrhizal with and beneficial to numerous pine and ñr species. Its epiphytic habit on stems does not damage

seedlings.

39
Deficiencies of ectomycorrhizal fungi in previously fumigated

nursery soils have been reported. Colonization of soil from airborne spores did not occur in these instances. Deficiencies could

be due to unfavorable weather conditions for mushroom production in forests adjacent to the nursery. Spores from these

mushrooms serve as inoculum for soil colonization. Also, the

nursery may be too far from forests harboring specific mushrooms. Research is currently underway to determine the feasibility and practical value of artificially infesting nursery soils

with specific ectomycorrhizal fungi known to be more beneficial

to tree survival and growth than those ectomycorrhizal fungi

which naturally occur on roots from airborne inoculum.

The consequence of soil fumigation to endomycorrhizal fungi

is another problem. Once these symbiotic fungi have been eradicated from soil, reinfestation is very slow because their spores

are not normally wind disseminated. Therefore, deficiencies of

these fungi in nursery soil following fumigation are not unusual. Endomycorrhizal development following successful soil

fumigation is from inoculum of the symbionts (1) still viable

in soil depths beyond effective fumigant penetration, (2) washed

in from water runoff or heavy rain splash from nonfumigated

areas of soil, (3) brought in by nonfumigated soil on cultivation

equipment, or (4) possibly from windblown nonfumigated soil.

Each of these possibilities would result in a slow development of

endomycorrhizae.

Considerable research has demonstrated that ectomycorrhizae

are not only beneficial to growth of trees, but are actually indispensable for survival and growth of transplant stock, in

particular Pinus spp. Thus, to insure survival and good growth

of normally ectomycorrhizal trees, seedlings used in reforestation

should have abundant ectomycorrhizae. It is not known if this

statement can be applied to endomycorrhizae. The significance

of endomycorrhizae to survival and growth of several species of

hardwood seedlings is currently being investigated.

Selected References

Clark, F. Bryan.

1969. Endotrophic mycorrhizal infection of tree seedlings with Endogone

spores. For. Sei. 15:134-137.

Hacskaylo, Edward.

1971. Mycorrhizae. U.S. Dep. Agrie. Misc. Publ. 1189, 255 p.

Jorgensen, J. R., and Eugene Shoulders.

1967. Mycorrhizal root development vital to survival of slash pine nursery stock. Tree Planters' Notes 18(2) :7-ll.

Kleinschmidt, G. D., and J. W. Gerdemann.

1972. Stunting of citrus seedlings in fumigated nursery soils related

to the absence of endomycorrhizae. Phytopathology 62:1447-1453.

Marx, Donald H., and W. Craig Bryan.

1971. Influence of ectomycorrhizae on survival and growth of aseptic

seedlings of loblolly pine at high temperature. For. Sei. 17:37-41.

Shoulders, Eugene, and J. R. Jorgensen.

1969. Mycorrhizae increase field survival of planted loblolly pine. Tree

Planters' Notes 20(1) : 14-17.

40
IL STEM AND BRANCH DISEASES (CANKERS)

The diseases of stems and branches are known as cankers,

diebacks, or galls, depending upon the symptoms they cause on

the host. Cankers are sunken areas of dead tissue in the bark,

while diebacks are the dying and/or curling of the shoot tip.

Galls are local enlarged or swollen portions of the branch or

stem. The causal fungi which may often fruit on the canker or

gall are helpful in diagnosing these diseases. Stem and branch

diseases are easily noticed in the beds when associated, as they

often are, with dead foliage. Without the dead foliage they may

go unnoticed until the grading and sorting operation. The following eight diseases are the most important and better-known

stem and branch diseases of forest tree seedlings.

41
11. PHOMOPSIS CANKER OF DOUGLAS-FIR

Diaporthe lokoyae Funk,

Phomopsis lokoyae Hahn

RICHARD S. SMITH, JR.

Phomopsis canker caused by Diaporthe lokoyae (imperfect

stage = Phomopsis lokoyae) is one of the most frequently encountered cankers of Douglas-fir in western North America. It is

a native disease which is most severe in the seedling and sapling

stages in natural stands, plantations, and forest nurseries.

Until recently only the imperfect stage, Phomopsis lokoyae, of

this fungus was known, and the disease was named after this

stage. In 1968, Funk described the perfect stage Diaporthe

lokoyae of this pathogen. Both stages of the fungus may occur

on the same specimen, but the fruiting bodies do not intermingle.

The perfect stage is usually on the distal portion of the killed

twig.

Distribution

Phomopsis canker was first described in 1933 from northern

California. It has since been found in the forests of Oregon,

Washington, and British Columbia, and has been reported as a

problem in some nurseries in these areas.

Hosts and Damage

Both the Pacific Coast and Rocky Mountain forms of Douglasfir are susceptible to the disease. Western hemlock and western

redcedar have been reported as rare hosts.

The disease causes cankers on the stems and branches. The

foliage and branch distal to the infection then yellows and dies

quickly (fig. 11-1). The disease kills some trees and causes

others to be forked and misshaped. The major loss results from

the culling of misshapen stock.

Life History

The sunken appearance of the canker that develops on young

stems and branches is due to continued diameter growth of

neighboring uninfected tissue while infected tissue has died.

Small, black, spherical fruiting bodies (pycnidia) develop in the

42
r-522511

Figure 11-1.—Douglas-flr seedling infected by Diaporthe lokogae.

cankered areas. These pycnidia produce spores which are waterborne, and spread to new hosts by the splashing drops of rain

or sprinkler irrigation. Under favorable conditions the spores

germinate and infect the new hosts. The perfect stage may develop later on the infected branch. This stage releases a windborne spore which is responsible for the long-distance spread of

the fungus.

In natural stands the intensity of infection of Phomopsis

canker varies greatly from year to year. Infection appears to

be more frequent when protracted rains are followed by periods

of drought. The intensity of this disease also fluctuates in nurseries from year to year, for reasons not yet explained.

43
Control

Periodic spraying with protective fungicides throughout the

growing season has effectively controlled this disease.

Selected References

Boyce, J. S.

1933. A canker of Douglas-fir associated with Phomopsis lokoyae. J. For.

31:664-672.

Boyce, John Shaw.

1961. Forest pathology. 3d Ed., 572 p. McGraw-Hill Book Co., New York.

Funk, A.

1968. Diaporthe lokoyae n. sp. the perfect state of Phomopsis lokoyae.

Can. J. Bot. 46:601-603.

44
12. SIROCOCCUS TIP BLIGHT

Sirococcus strobilinus Preuss.

RICHARD S. SMITH, JR.

Sirococcus tip blight is a disease affecting- chiefly the newshoots of conifers. It is caused by a fungus producing only waterborne (spread) spores, and is also referred to by the name

Ascochyta piniperda.

Distribution

This disease occurs in both forests and forest nurseries in

Europe, and in eastern North America. Recently it has been

found in natural reproduction in the Lakes States, Alaska, and

British Columbia. In California, it is restricted to the north

coastal fog belt.

Hosts and Damage

S. strobilinus has been reported in the United States and

Canada on seedlings of Norway and red spruces. Coulter, Jeffrey,

sugar, red, lodgepole, and ponderosa pines, and western hemlock.

The disease causes a tip dieback and stem and branch cankers

in the current year's growth. The foliage distal to the infection

becomes yellow and dies. This disease kills trees with multiple

infections, and causes others to become forked and misshapen.

Nursery losses are due both to tree death and to culling of misshapen stock.

Life History

Initial infection occurs in the juvenile needles. The fungus

spreads down into the succulent stem tissues, where it causes

a small purplish canker (fig. 12-1). The infection enlarges

longitudinally more rapidly than around the stem, producing an

elongate, sunken canker. A small drop of resin is often exuded

F-522512

Figure 12-1.—A Coulter pine seedling

infected with Sirococcus strobilinus. Curling of the top is typical

of this disease.

45
in the center of the canker at the needle base from which stem

infection originated. When infection occurs in the region of

elongation, the restricted growth in the cankered area causes

the shoot tip to curl over and form a crook (fig. 12-2). As the

canker matures, small, black fruiting bodies begin to appear on

the infected needles and the older portions of the canker (fig.

12-3). These fruiting bodies produce large masses of spores

which are spread to new hosts by the splashing of water drops

from rain or sprinkler irrigation. Under favorable conditions of

high humidity and mild temperatures (above 10° C), the spores

germinate and infect the new host.

F-522513

Figure 12-2.—A young canker caused

by Sirococcus strobilinus developing at the base of an infected juvenile needle of a 2-year-old Jeffrey

pine.

Figure 12-3.—The small, black fruiting bodies of Sirococcus strobilinus

developing in an older cankered

area.

Control

Periodic spraying with protective fungicides throughout the

growing season will effectively control this disease. Chlorothalonil (Bravo W-75 and 6F) has successfully controlled this disease in West coast conifers.

Selected References

Funk, A.

1972. Sirococcus shoot-blight of western hemlock in British Columbia and

Alaska. Plant Dis. Rep. 56:645-648.

Graves, Arthur H.

1914. Notes on diseases of trees in the southern Appalachians. Phytopathology 4:63-72.

Smith, R. S., Jr., A. H. McCain, M. Srago, R. F. Krohn, and D. Perry.

1972. Control of Sirococcus tip blight of Jeffrey pine seedlings. Plant

Dis. Rep. 56:241-242.

Smith, R. S., Jr.

1973. Sirococcus tip dieback of Piniis spp. in California forest nurseries.

Plant Dis. Rep. 67:69-73.

46
13. GREY MOLD OF GIANT SEQUOIA

Botrytis cinérea (Fr.) Pers.

RICHARD S. SMITH, JR.

Grey mold—a disease of many plants, including forest tree

seedlings—is caused by the fungus Botrytis cinérea; its perfect

stage is Sclerotinia fuckeliana (deBy) Fckl. This fungus, also

a saprophyte, is almost universally present on dead and dying

vegetable matter. As a plant pathogen, the fungus usually must

first become established on dead or moribund parts of a host

F-62251B

Figure 13-1.—A 1-year-old giant sequoia with a stem canker caused by

Botrytis cinérea.

47
plant. It then can spread into adjacent healthy tissues. A few

species of conifers, particularly giant sequoia, are so susceptible

to B. cinérea that the fungus can successfully infect healthy

green tissues.

Distribution

Since the fungus is found in all temperate regions, the presence of the disease is dependent only on having a susceptible

host under suitable conditions. It has been reported on various

forest species from Asia, Europe, and North America.

Hosts and Damage

This pathogen has an extremely large host range. Nearly all

forest trees are susceptible to some degree, but it is a serious

problem only on a fev^ species such as redv^ood, giant sequoia,

and Monterey and Italian cypress.

The disease causes a branch dieback and canker of green

succulent tissues. In 1-year-old seedlings, stem cankers originating from infected lower branches frequently girdle the stem

(fig. 13-1) and kill the entire seedling. In California, when

conditions are suitable, whole sections of nursery beds of giant

sequoia have been destroyed.

Life History

Under prolonged periods of high humidity and cool temperatures, this fungus infects the juvenile bracts and young branches,

particularly the lower shaded branches. Once established, it

proceeds downward into the stem, killing the tissues it infects.

In the succulent 1-year-old stem tissues, it forms a black sunken

canker which eventually girdles the stem. The portion of the

seedling distal to the girdle dies. Since the lower shaded branches

are infected more often than the upper branches, the girdles are

usually low on the stem in a position where most, or all, of the

crown is killed. This disease is favored by cultural practices such

as high planting densities or lath shading, which limit air movement and raise the humidity around the seedling.

Under moist conditions, B. cinérea can be seen as a thin grey

web of mycelium on infected plant parts. From this mycelium

and from infected tissue, tufts of black conidiophores arise bearing clusters of white to grey spores. When the conidiophores are

brushed, clouds of spores are released.

Control

Proper cultural practices which increase aeration and decrease

humidity will help reduce or prevent losses in most species. In

highly susceptible species, fungicidal sprays may be required.

48
Most fungicides act as protectants ; they do not cure the disease

once it is in the plant. Therefore, fungicides must be applied at

the first sign of the disease, and their use continued as long as

conditions favorable to the disease persist.

Grey mold is also a serious problem during the cold storage

and transportation of forest seedlings. Whole shipments of

seedlings have been lost to this disease. To reduce chances of

losses, all seedlings infected v^ith grey mold must be culled out at

the time of sorting and packaging. The seedlings should be

protected further by a fungicide spray or dip just before packaging.

Selected References

Halber, Max.

1963. Botrytis sp. on Douglas-fir seedlings. Plant Dis. Rep. 47:556.

Peace, Thomas Ronald.

1962. Pathology of trees and shrubs. 753 p. Oxford Univ. Press, London.

Smith, Richard S., A. H. McCain, and M. D. Srago.

1973. Control of Botrytis storage rot of giant sequoia seedlings. Plant

Dis. Rep. 57:67-69.

49
14. SCLERODERRIS CANKER

Scleroderris lagerbergii Gremmen

DARROLL D. SKILLING

Scleroderris canker kills both seedlings and plantation trees.

Mortality is most rapid among nursery seedlings; stems are

killed by girdling. Planting infected seedlings is especially

serious, as the infected trees provide inoculum for future disease

buildup.

It is not known if Scleroderris canker is native to the United

States, but it has been present for many years. The fungus responsible for this disease was not identified until 1964 in the

United States. It is currently causing heavy losses in red and

jack pine plantations in northern Wisconsin and Michigan.

Distribution

The disease is known to be present in Michigan, Wisconsin,

Minnesota, Vermont, and New York. It is also widely distributed

throughout Ontario and Quebec, and occurs in New Brunswick

and Nova Scotia. Short growing seasons, with attendant prolonged damp springs and autumns, and heavy snowfall seem to

favor disease development.

Hosts and Damage

Scleroderris canker has been found on Scotch, red, and jack

pines in the United States. In addition, white and lodgepole

pines and white and black spruces have been infected in Quebec.

Austrian, Corsican, Swiss Mountain, Swiss stone, and Maritime

pines are known hosts in Europe, as well as Norway spruce and

Douglas-fir, and Todo fir in Japan.

Damage to pines includes death of lateral and terminal

branches. Infection is usually limited to the lower 6 feet of the

tree. The fungus commonly grows into the main stem from

infected branches, resulting in a stem canker. A characteristic

yellow-green discoloration is frequently observed in the cambial

zone of recently killed tissue. Infected needles on red pine

nursery stock are orange at their base during early May. These

needles are loose and can easily be removed from the seedling

(fig. 14-1).

In Quebec the symptoms of Scleroderris infection on black

spruce include dead or dying stems, leaders, and branches.

50
F-622516

Figure 14-1.—Red pine seedlings (3-0) infected with Scleroderris lagerbergii.

The orange discoloration at the base of the needles is obvious during early

spring in North America.

Life History

Primary infection is by windblown ascospores disseminated

during moist weather from May to October, although most are

released during June and July. Infection is through buds or

needles. Infected branch tips are usually dead the following

summer. The fungus grows down the branch until it reaches

the main stem of the tree. On trees less than 4 years old the

stem is quickly girdled, and the tree is dead within a few months.

A few months after a branch dies, pycnidia appear near the

base of dead needle fascicles. Asexual spores ooze out of these

pycnidia during wet weather. These spores are transported by

rain splash to nearby branches, which may become infected.

Apothecial fruiting bodies appear in early summer on branches

that have been dead for 1 or 2 years. The apothecia are also

usually found at the base of dead needle fascicles, and are often

closely associated with the pycnidial stage. Ascospores become

mature in the apothecia by late May, and are discharged following rainfall from then until October.

Control

Infection by S". lagerbergii can be prevented in nursery beds

by applying approximately seven sprays of chlorothalonil

(Bravo W-75 and 6F) at 3 pounds of 75 percent wettable

powder or II4 quarts of 6-pound a.i. liquid formulation per 100

51
gallons of water. Applications should begin as soon as new shoot

growth starts. They should be repeated at 2-week intervals until

the first of July, then at 4-week intervals until the first of September. One or two extra sprays may be needed if rainfall is

unusually heavy during June.

Selected References

Dorworth, C. E.

1970. Scleroderris canker in Ontario forest nurseries. For. Res. Lab.,

Sault Ste. Marie, Ont. Inf. Rep. O-X-148, 9 p.

Ohman, John H.

1966. Scleroderris lagerhergii Gremmen: The cause of dieback and mortality of red and jack pines in Upper Michigan plantations. Plant

Dis. Rep. 50:402-405.

Skilling, Darroll D., and Charles E. Cordell.

1966. Scleroderris canker on National Forests in Upper Michigan and

northern Wisconsin. USDA For. Serv. Res. Pap. NC-3, 10 p. N.

Cent. For. Exp. Stn., St. Paul, Minn.

Skilling, Darroll D., and Clinton D. Waddell.

1970. Control of Scleroderris canker by fungicide sprays. Plant Dis.

Rep. 54:663-665.

Smerlis, E.

1967. Occurrence and pathogenicity of Scleroderris lagerhergii in Quebec.

Plant Dis. Rep. 51:584-585.

52
15. SOUTHERN FUSIFORM RUST

Cronartium fusiforme Hedge. & Hunt Ex Cumm,

S. J. ROWAN*

Fusiform rust is the most serious nursery disease of slash and

loblolly pines in the southern United States. Without fungicida!

control, this disease may affect up to 90 percent of all slash and

loblolly pine seedlings in a given nursery. Although fungicidal

sprays are routinely used in southern nurseries, in most years

up to 20 percent of the slash and loblolly pine nursery stock is

infected in high-hazard sites by this pathogen. Because infected

nursery seedlings rarely survive the first year of outplanting,

nursery infection is essentially equivalent to seeding mortality.

r-622517

Figure 15-1.—Typical galls on loblolly pine seedlings caused by Cronartium

fusiforme.

*Plant Pathologist, Forestry Sciences Laboratory, Southeastern Forest

Experiment Station, USDA Forest Service, Athens, Ga.

53
Distribution

Cronartium fusiforme is indigenous to the southern United

States from Maryland to Florida and westward to Arkansas

and Texas. The incidence of the disease is highest in a zone

approximately 150 miles wide extending from coastal South

Carolina to Texas.

Hosts and Damage

Natural and artificial inoculations have proved 32 species of

pines to be susceptible to the disease. Slash, loblolly, and South

Florida slash pines are highly susceptible, longleaf and pond

pines are intermediate, and pitch and shortleaf pines are resistant.

Natural and artificial inoculations have also proved 33 species

of oak to be susceptible to the disease. Four species of chestnut, two species of chinkapin, and tanoak have also been shown

to be susceptible, but members of the black oak group are the

most common alternate hosts of the fungus.

Typically spindle-shaped stem and branch galls result from

infections of pine (fig. 15-1). Seedlings infected in the early

stages of growth often die when outplanted whereas others

develop bushy crowns because of the loss of apical dominance. The

reduced mechanical strength of infected stem tissue of older

r-522518

Figure 15-2.—Telia on the underside of water oak leaves caused by Cronartium.

fusiforme.

54
pines results in increased wind breakage. Infections of the alternate (oak) hosts generally cause minor leafspot injury; in

unusually heavy infections, there may be some defoliation.

Life History

Aeciospores, produced on pine galls in early spring, infect expanding oak leaves. Uredial pustules develop rapidly on infected

oak leaves and produce uredospores v^hich may infect other

young leaves. As these infected leaves mature, telia appear and

produce basidiospores which infect new pine needles and

shoots (fig. 15-2). The basidiospores are produced from spring

to early summer during periods of high humidity and moderate

temperatures. On nonhost tissue, the basidiospores may form

secondary spores which eventually can germinate directly on

pine tissue and cause infection.

Control

Fusiform rust infections can be prevented by spray applications of ferbam or ziram. Ferbam is the most commonly used

fungicide. A spreader-sticker is usually added to improve coverage and reduce weathering. The frequency and coverage of

fungicidal sprays is important for effective control. During the

infection season, sprays are usually applied twice each week and

after rains. Sprays applied before hazardous rust weather is

forecast are more effective than those applied after such periods.

All infected seedlings should be culled and only healthy seedlings

outplanted.

Selected References

Czabator, Felix J.

1971. Fusiform rust of southern pines—a critical review. USDA For.

Serv. Res. Pap. SO-65, 39 p. South. For. Exp. Stn., New Orleans,

La.

Davis, R. T., and G. A. Snow.

1971. Forecasting weather favorable for fusiform rust infection. Tree

Planters' Notes 22(2) :3-4.

Foster, A. A.

1959. Nursery diseases of southern pines. U.S. Dep. Agrie. For. Pest

Leafl. 32, 7 p.

Hedgecock, George G., and Paul V. Siggers.

1949. A comparison of the pineoak rusts. U.S. Dep. Agrie. Tech. Bull.

987, 30 p.

Hodges, Charles S., Jr.

1962. Diseases in southeastern forest nurseries and their control. U.S.

Dep. Agrie, For. Serv., Southeast. For Exp. Stn., Stn Pap. 142, 16 p.

Hodges, Charles S., Jr., and John L. Ruehle.

1969. Nursery diseases of southern pines. U.S. Dep. Agrie. For. Pest

Leafl. 32, 8 p.

Siggers, Paul V.

1955. Control of the fusiform rust of southern pines. J. For. 53:442-446.

55
16. WESTERN GALL RUST

Peridermium harknessii J. P. Moore

RICHARD S. SMITH, JR.

Western gall rust, although often found in the forests of

western North America, has been only an occasional problem in

forest nurseries. Although not generally a problem to nursery

production of pines, western gall rust can have serious consequences when outplanted. Not only are infected trees doomed,

but the rust could easily spread from the seedlings to the surrounding forest.

Distribution

This disease is found generally throughout the pine forests of

western North America, and in scattered locations in eastern

United States and Canada. In western forest nurseries the infection has been limited to a few nurseries with a natural source

of inoculum nearby.

Hosts and Damage

The western gall rusts caused by the fungus Peridermium

harknessii (sometimes called Endocrormrtium harknessii) y attacks a large number of native hard pines, including knobcone,

jack, lodgepole, Coulter, Jeffrey, bishop, ponderosa, Monterey,

and Digger pines as well as the exotic Canary Island, Aleppo,

Swiss Mountain and Scotch pines.

This disease causes globose galls or swellings at the site of

infection (fig. 16-1). Gall rust may kill the entire seedling, or

just that portion distal to the infection. The seedling may live

and grow, but as the gall enlarges, the stem becomes misshapened and weakened (fig. 16-2). Excess branching and loss

of apical dominance may result in "brooming'' of the tree at

the site of infection. Such damage is not usually seen in the

nursery, but develops after the seedling is outplanted. Nevertheless, damage is all directly attributable to the disease contracted in the nursery; hence, all gall-bearing seedlings should

be culled.

56
r-B22519

Figure 16-1.—A lodgepole pine seedling infected with western gall rust

shows a globose gall and the orange

spore masses in the bark cracks.

F-522520

Figure 16-2.—Lodgepole pine seedlings infected with western gall

rust.

Life History

Because it takes 2 to 4 years before an infection starts producing spores, the source of infecting spores is usually outside

the nursery beds. The source may be infected pines in ornamental plantings, in windbreaks, or in surrounding pine forests.

In spring and early summer, orange-colored aeciospores are

produced on infected trees and dispersed by wind. Under the

right temperature and moisture conditions, the spores deposited

on young shoots of a susceptible pine germinate and infect the

seedling. Either late the same year or during the next growing

season, a globose gall or swelling begins to develop at the site

of infection. In spring or early summer of second or third year,

orange-colored masses of aeciospores are produced on these

galls coloring them a distinctive bright orange. The aeciospores

are formed within thin white sacs underneath and between the

bark scales. When the sacs mature, they rupture and release

aeciospores into the wind.

Control

The most effective and economical method of control is to remove the source of infecting spores. All gall-bearing pines within

a half mile of the nursery should either be pruned free of all

galls or felled. These sanitation measures have reduced the

amount of western gall rust in the nurseries in which the disease was a problem.

57
All gall-bearing seedlings should be destroyed. Because there

are many latent infections with this disease, it is also desirable

to check all plantations in which stock had been exposed to infection. Also, care should be exercised in the movement of hard

pine nursery stock from presently infested areas into those

areas which are presently free of this disease.

Selected References

Boyce, John Shaw.

1961. Forest pathology. 3d Ed., 572 p. McGraw-Hill Book Co., N.Y.,

Toronto, London.

Merrill, W.

1972. Occurrence of Endocronartium harknessii in Pennsylvania. Plant

Dis. Rep. 56:1058.

Peterson, Glenn W.

1973. Dispersal of aeciospores of Peridermium harknessii in Central

Nebraska. Phytopathology 63:170-172.

Peterson, Roger S.

1960. Western gall rust on hard pines. U.S. Dep. Agrie. For. Pest Leafl.

50, 8 p.

Peterson, Roger S.

1967. The Peridermium species on pine stems. Bull. Torrey Bot. Club

94:511-542.

58
17. JACK PINE RUSTS

Cronartium sp.,

Peridermium sp.

DARROLL D. SKILLING

Jack pine seedlings are attacked by a number of stem rust

fungi that frequently cause much mortality. These fungi include Cronartium quercuum, eastern gall rust; C. comptoniae,

sweetfern rust, C. comandrae, comandra rust; C. coleosporioides,

stalactiform rust; and Peridermium harknessii, western gall

rust. With the exception of P. harknessii, which is discussed in

article 16 of this Handbook, all of these rust fungi require an

alternate host to complete their life cycle. Seedling losses often

occur where an alternate host of these rusts is present near jack

pine nurseries.

Distribution

Eastern gall rust is found east of the Great Plains from Canada to southern United States. Sweetfern rust is transcontinental in distribution from Nova Scotia west to British Columbia and Alaska, in the eastern United States south to North

Carolina, and in the western United States in Washington and

Oregon. Comandra rust occurs from New Brunswick to the

rAV/'

F-522521

Figure 17-1.—Jack pine stem rusts on seedlings: Left to riglit, eastern gall,

comandra, sweetfern.

69
Yukon and British Columbia in Canada and southward to California, New Mexico, and Alabama in the United States. It is

present in all States west of the Rocky Mountains. Stalactiiorm rust is present from New Brunswick to British Columbia

and south to California, central Colorado, Minnesota, and

Michigan.

Hosts and Damage

Disease

Eastern gall rust

Sweetfern rust

Comandra rust

Stalactiform rust

Aecial Host

Jack, Scotch, shortleaf,

Virginia, and other hard

pines

Jack, Scotch, lodgepole,

ponderosa, pitch, and

other hard pines

Jack, Scotch, lodgepole,

ponderosa, loblolly,

shortleaf, and other pines

Jack, lodgepole, Jeffrey

and ponderosa pines

Uredial and Telial Host

Various species of oaks

Sweetfern and sweetgale

Bastard toad flax and

northern comandra

Cow-wheat, Indian paintbrush, lousewort, and

yellow-rattle

Eastern gall rust produces globose-shaped galls on the stems

oí young pine seedlings. In problem areas, nursery losses frequently exceed 25 percent. Infected seedlings that are outplanted

are subject to wind breakage in later years, because the galls

greatly weaken the stem, or result in considerable cull

Sweetfern rust usually is evident as an elongate swelling on

the stems of infected seedlings (fig. 17-1). This swelling appears 1 to 2 years following infection. The disease is most conspicuous m early spring when orange-yellow aeciospores are

produced on the gall. Nursery seedlings grown near alternate

host plants are often heavily damaged by the girdling action of

this rust In one Michigan nursery, sweetfern rust destroyed

nearly all the lodgepole and ponderosa pine seedlings.

The first indication of comandra rust on seedlings is a spindleshaped swelling of the bark near the base of the stem. Cankers

originate on needle-bearing twigs and stems. Cankered bark

on which aecia are produced, is cracked and pitted. In time

branches and stems are girdled. Mortality, however, is usually

not excessive.

The spindle-shaped cankers on seedlings infected with stalactiform rust are not easily distinguishable from those produced by

sweetfern and comandra rusts. Elongate diamond-shaped cankers are produced on mature trees. Ponderosa pine nursery stock

has been severely damaged by stalactiform rust in Montana.

Life History

The biological development of the four rusts reported here is

similar All require an alternate host to complete their life

cycle. In each case this is a broadleaf plant entirely unrelated

60
to the pine host. A typical life cycle starts with a basidiospore

from the alternate host germinating on a pine needle. The rust

fungus penetrates the needle and grows down into the stem of

the seedling. During the summer, 1 to 3 years after infection,

drops of a thick, somewhat sticky and reddish-orange liquid

begin to exude from the diseased bark. These drops contain

pycniospores that probably function in the sexual phase of reproduction of the fungus. Approximately 1 year later, orangecolored aeciospores are produced in the same area as the

pycniospores.

The wind-disseminated aeciospores can infect the alternate

host under proper temperature and moisture conditions. About

2 weeks after the alternate host is infected, orange-colored

urediospores are produced on the undersurface of the leaves.

These urediospores can infect additional alternate host plants,

increasing the disease inoculum in the area. In late summer or

early fall, brownish, hairlike structures called telia are produced on the infected leaves. In the case of eastern gall rust this

may occur in early spring. Teliospores germinate under cool,

moist conditions and produce windblown basidiospores which

again infect the pine host.

The two critical periods in the development of rust epidemics

are the period of aeciospore dissemination to the alternate host,

and the period when basidiospores are formed and infect the

pine seedlings. Long periods of moist weather during these spore

discharge periods are conducive to severe infection.

Control

Control measures may be used to reduce jack pine stem rust

losses. The weak link in the life cycle of these rusts is their

need to live for a time on a broadleaf host. Nurserymen can

take advantage of this weakness by removing the broadleaf

hosts from the nursery area whenever practical. The exact distance necessary to eradicate alternate hosts is not known, but is

probably from 14 to 1 mile.

Infected seedlings should be eliminated at the nursery. Planting infected seedlings may allow the fungus to spread to the

alternate host and back to other pines in the plantation on

nearby forests, creating an epidemic.

Selected References

Anderson, Gerald W.

1963. Sweetfern rust on hard pines. U.S. Dep. Agrie. For. Pest Leafl. 79,

7 p.

Anderson, Neil A.

1963. Eastern gall rust. U.S. Dep. Agrie. For. Pest Leafl. 80, 4 p.

Anderson, Neil A., David W. Freneh, and Ralph L. Anderson.

1967. The stalaetiform rust on jaek pine. J. For. 65:398-402.

Mielke, J. L., R. G. Krebill, and H. R. Powers, Jr.

1968. Comandra blister rust of hard pines. U.S. Dep. Agrie. For. Pest

Leafl. 62, 8 p.

61
18. WHITE PINE BLISTER RUST

Cronartium rihicola J. C. Fisher

DARROLL D. SKILLING

White pine blister rust, caused by the fungus Cronartium ribi^

cola, was introduced into North America from Europe about

1900. Economically, it is one of the most important forest diseases m the United States and Canada; it causes an annual

growth loss and mortality in excess of 200 million cubic feet.

In many areas of North America where conditions for blister

rust development are favorable, it is responsible for the virtual

abandonment of susceptible pine species from forest planting

programs.

Distribution

White pine blister rust is distributed throughout most of the

range of white pines in North America. In Canada the disease

IS present m British Columbia, Ontario, Quebec, and portions

of Newfoundland. In the western United States it is present

throughout Washington, Oregon, northern California, northern

Idaho, Montana, and Wyoming. In the eastern United States it

is present in the New England States west to Minnesota and

south into the highlands of North Carolina.

Hosts and Damage

The fungus responsible for the blister rust disease spends

part of its life cycle on the five-needled pines and the remainder

on plants m the genus Ribes (currants and gooseberries).

The most susceptible pines are western white, sugar, limber,

and white-barked pines. Eastern white pine and foxtail pine are

moderately susceptible. Swiss stone pine and Balkan pine, both

native to Europe and Asia, are quite resistant. Pinyon pine and

Oriental white pine are not susceptible to the blister rust fungus.

All of the native North American Ribes are susceptible to

blister rust. The European black currant, Ribes nigrum, is

highly susceptible. In the white pine regions of Idaho, A. petio-

¿are and Ä. merme are most susceptible ; Ä. viscosissium is moderately susceptible and R. lacustre is the least susceptible.

62
White pine blister rust is evident as an elongate swelling on

the stems of infected nursery seedlings. This swelling frequently becomes yellow orange to copper 1 year after infection.

The disease is most conspicuous in early spring when orangeyellow aeciospores are produced on the swollen area. White pine

seedlings grown near large concentrations of Ribes plants are

often heavily infected by C. ribicola. Infected stock is frequently shipped to the field because new infections are difficult

to recognize.

Blister rust kills white pines of all ages, with the smaller

trees dying more rapidly. The fungus grows directly from infected needles into the main stem of nursery seedlings. Stem

girdling kills the seedlings. Branches girdled by the fungus on

older trees develop brown needles called "flags" which can be

easily recognized. The fungus frequently grows into the main

stem from these infected branches, resulting in stem cankers.

These cankers may persist for years on larger trees, and eventually kill the tree by girdling the stem.

F-622B22

Figure 18-1.—White pine blister rust canker on young pine. The creamcolored blisters contain the aeciospores that infect Ribes.

63
Life History

The life cycle starts with a basidiospore from an infected

Rihes plant germinating on a pine needle during late summer or

early fall. The rust fungus penetrates into the needle and grows

down into the stem of the seedling. This mycelium continues to

advance as long as the host remains alive. During the early

summer, 2 to 4 years after infection, yellow-brown blisters appear on the bark and drops of a sticky reddish-orange liquid

ooze out. These drops contain pycniospores that probably function m the sexual phase of reproduction of the fungus. Approximately 1 year later, aeciospores are produced in the same areas

as the pycniospores. Cream-colored blisters appear which burst

and release the orange-colored aeciospores (fig. 18-1). The

aeciospores are carried by the wind, sometimes for several hundred miles, to infect Ribes leaves.

About 2 weeks after infection, orange-colored urediospores

are produced on the undersurface of the Rihes leaves. These

urediospores can infect additional Rihes leaves, which increases

the disease inoculum in the area. Up to seven generations of

urediospores may be formed in a season. In late summer or

early fall, brownish, hairlike structures called telia are produced on the infected leaves. Teliospores are produced on the

tehal columns. These germinate and produce basidiospores,

which are blown to the needles of pine to complete the life cycle.

Moisture and cool temperatures (15° C is optimum for spore

germination) favor infection of both hosts. Basidiospores are

produced only when the relative humidity is high, especially

after rains and during fogs. The basidiospores are quite delicate, and if exposed to dry air or sunlight they die quickly. The

effective range over which pines can be infected by basidiospores

IS usually a few hundred feet. However, under ideal conditions

of cool, moist weather, it is possible for basidiospores to infect

pines over a mile away.

Control

White pine blister rust can be controlled by removing the

Rihes bushes within 14 to 1 mile of the nursery. This method of

control is possible because the fungus cannot spread from pine

to pine, but must live for a time on the Rihes plant to complete

its life cycle. When possible, white pine seedlings should be

grown only in nurseries where the environment is unfavorable

for blister rust infection because of high temperatures and low

humidity.

Infected seedlings should be culled at the nursery whenever

disease symptoms appear. The planting of infected seedlings

may allow the fungus to spread to Rihes plants and back to

other pmes m the plantation, creating an epidemic.

The most promising control for blister rust is the selection

and breeding of resistant trees. Substantial progress has been

64
made in this area in the last 10 years. Seed orchards are now

being established with trees that have shown resistance to C.

ribicola.

Selected References

Benedict, W. V.

1967. White pine blister rust. p. 184-188. In Important Forest Insects

and Diseases of Mutual Concern to Canada, the United States, and

Mexico. Can. Dep. For. Rural Develop., Publ. 1180, 248 p. Ottawa.

Peace, Thomas Ronald.

1962. Pathology of trees and shrubs. 753 p. Oxford Univ. Press, London.

Spaulding, Perley.

1922. Investigations of the white pine blister rust. U.S. Dep. Agrie.

Bull. 957, 100 p.

Van Arsdel, E. P.

1964. Growing white pines to avoid blister rust—new information for

1964. U.S. For. Serv. Res. Note LS-42, 4 p. Lake States For. Exp.

Stn., St. Paul, Minn.

65
III. FOLIAGE DISEASES

Foliage diseases attack leaves, needles, and young shoots. They

are called leaf spots when only a portion of the leaf is affected,

and leaf and needle blights when the whole leaf or needle is

killed. Anthracnoses are foliage diseases caused by a particular

group of fungi in which both the leaves and young developing

shoots are killed. In general, foliage diseases are more easily

recognized in early stages and easier to control than are the

root and canker diseases. The following diseases are the most

important and common foliage diseases of forest tree seedlings

in North America.

66
19. LOPHODERMIUM NEEDLE CAST OF PINES

Lophodermium pinastri (Schrad.) Chev.

DARROLL D. SKILLING

Although long considered a serious disease in European nurseries and plantations, Lophodermium pinastri has only recently received attention as a major disease in the United

States. It is currently causing serious losses in red and Scotch

pine nurseries in the eastern United States.

Distribution

The disease occurs throughout the United States. Severe

damage to nursery stock has been reported in Michigan, Wisconsin, Minnesota, West Virginia, Washington, Massachusetts,

Pennsylvania, North Carolina, South Carolina, and Nova Scotia.

Hosts and Damage

L. pinastri has a wide host range. It is known to attack at

least 26 species and varieties of pines. In North America the

F-B22623

Figure 19-1.—Red pine seedlings (2-0) infected by Lophodermium pinastri.

Green trees in foreground were protected with the fungicide maneb.

67
most serious damage has been to red and Scotch pines. Other

pme hosts include eastern white, jack, ponderosa, Mugo, Austrian, Jeffrey, western white, Monterey, lodgepole, shortleaf,

pitch, loblolly, longleaf, and slash.

The first symptoms of the disease are brown-black spots on

mfected needles. Greenish-brown bands appear on infected

needles in late summer and fall. These infected needles turn

brown the following spring. On most pine hosts the damage

from L. pinastri has been limited to premature dropping of

only part of the needles. In many forest nurseries, however, entire beds of red and Scotch pine seedlings have been killed by

extensive needle defoliation or terminal bud dieback (fig. 19-1).

Smce 1966, several million trees 1 to 4 years old have, been

killed by this disease. Mortality has been especially severe in

1- and 2-year-old trees. In addition, the quality and vigor of

millions of surviving trees have been reduced by severe needle

loss. Heavy losses in field plantations have resulted when seedImgs were outplanted before it was apparent that they were

mfected. In many cases, symptoms developed after field planting.

Life History

Infection appears to be entirely by airborne ascospores, which

germmate on the pine needles. The spores are produced in tiny,

black, elongate fruiting bodies called hysterothecia. These develop on either attached or fallen needles during the late summer or fall following infection. During wet weather, the ascospores are forcibly discharged into the air. The major infection

period is from August to October.

Nursery infection may develop from either infected needles

in the seedbeds or from nearby infected windbreaks.

Control

Infection by L. pinastri can be prevented in nursery beds by

repeated applications of maneb or chlorothalonil (Bravo W-75

or 6F). Normally, four to six applications from mid-July to

September will give adequate protection.

Selected References

Boyce, John S., Jr.

1951. Lophodermium pinastri and needle browning of southern nines

J. For. 49:20-24.

NichoIIs, Thomas H.

1973. Fungicide control of Lophodermium pinastri on red pine nursery

seedlings. Plant Dis. Rep. 57:263-266.

NichoIIs, Thomas H., and Darroll D. Skilling.

1970. Lophodermium pinastri outbreak in Lake States forest nurseries

Plant Dis. Rep. 54:731-733.

Peace, Thomas Donald.

1962. Pathology of trees and shrubs. 753 p. Oxford Univ. Press, London.

68
20. BROWN SPOT NEEDLE BLIGHT

Scirrhia acicola (Dearn.) Siggers,

Systremma acicola (Dearn.) Wolf and Barbour

ALBERT G. KAIS*

Brown spot needle blight, the most serious disease of longleaf pine, both in the forest and in the nursery (fig. 20-1) has

been a primary reason for the widespread planting of other

pine species on former longleaf sites. Its effect on growth and

vigor in the nursery is severe, and a control program is essential.

F-622524

Figure 20-1.—Longleaf pine seedlings infected with brown spot.

Distribution and Hosts

Brown spot needle blight is common in nurseries in the natural range of longleaf pine (U.S. Coastal Plain from North

Carolina to Texas). It may infect loblolly and slash pines within

*Plant Pathologist, Southern Forest Experiment Station, USDA Forest

Service, Gulfport, Miss.
or beyond the range of longleaf. It is occasionally found on

other southern pines, and on some exotic nursery stock in the

south. Recently, it was reported on white pine nursery stock

in Tennessee.

The range of Scirrhia acicola is by no means restricted to the

south. The fungus has been reported in Missouri and Ohio, and

has recently appeared in jack pine plantations in Canada. It has

also caused severe damage in Scotch pine Christmas tree plantations in six North and Central States. Only the conidia! (imperfect) state of the fungus, Lecanosticta acicola (Thum.) Syd.,

has been found in northern areas.

Symptoms

Brown spot lesions may develop on secondary needles at any

season of the year, but appear most commonly from May

through October. The first evidence of infection is the presence

of small, circular, grayish-green spots. These become straw

yellow and later change to a light brown. A darker chestnutbrown discoloration often borders the spot when fruiting bodies

form in the lesion. The individual spots, which are about oneeighth inch in diameter, frequently coalesce. Eventually the

needle tissue beyond and between groups of spots dies. Needles

with multiple infections take on a mottled appearance. Another

type of lesion, known as bar spot, consists of an amber-yellow

band with a small brown center encircling the needle. Both

types of spots have well-defined margins.

The infected needle usually has three definite zones: (1) the

basal portion, which is green; (2) the middle portion, which is

spotted with lesions that alternate with green needle tissue;

and (3) the apical portion, which consists of dead needle tissue. The loss of needle tissue stunts the growth of nursery

seedlings and may kill them. If badly infected seedlings are outplanted they may be too weak to survive competition and dry

weather, or may remain in the grass stage for many years.

Life Cycle

The macroscopic fruiting bodies (acervuli), which are found

on lesions, produce conidia which rupture the leaf surface. The

conidia are cylindrical, curved, 1-4 septate, and olive green to

brown in color; they are exuded in a sticky mass which may

form a ribbon up to 1 mm in length. Conidia are disseminated

for short distances by rain splash and cause local intensification

of the disease.

In 2 to 3 months after initial infection, ascospores are produced in perithecia that are embedded within dead leaf tissue.

They are hyaline, oblong-cuneate, unequally bicellular, and have

two prominent brown oil drops in each cell. Mature ascospores

are discharged during periods of rain, heavy dew, and fog.

They are disseminated by wind. It is thought that these spores

70
initiate light, scattered infections, sometimes at great distances from their source, and are the principal means by which

the fungus invades nursery beds.

Control

Control in nurseries can be achieved with chemical sprays. A

freshly made 4-4-50 bordeaux mixture (4 pounds copper sulfate, 4 pounds hydrated lime, and 50 gallons of water) is economical and effective. No spreader-sticker is required. This mixture is applied at the rate of 60 gallons per acre at 300 pounds

pressure. Seedlings should be sprayed at frequent enough intervals to protect new foliage as it develops. Four to seven applications of bordeaux mixture from the end of May to October

are usually sufficient. Nursery managers in high-hazard areas

probably should spray every 2 weeks throughout the infection

period, and should schedule a final spraying just prior to lifting.

Maneb and chlorothalonil (Bravo) have been registered for

use on brown spot needle blight. Applications of maneb or

chlorothalonil should begin in spring or early summer, before

infection occurs, and should be repeated at 2-week intervals

through the end of summer. Extra applications should be made

after heavy rains. Sporulation and dissemination of the fungus

occurs in the deep South during unseasonably warm periods in

the winter months. Therefore, seedlings that are held beyond

the usual lifting date may benefit from a precautionary spray.

Recent research has revealed the existence of heritable resistance to brown spot needle blight, and resistant longleaf nursery stock may thus become available.

Selected References

Lightle, Paul C.

1960. Brown-spot needle blight on longleaf pine. U.S. Dep. Agrie. For.

Pest Leafl. 44, 7 p.

Siggers, Paul V.

1944. The brown spot needle blight of pine seedlings. U.S. Dep. Agrie.

Tech. Bull. 870, 36 p.

Wakeley, Philip C.

1954. Planting the southern pines. U.S. Dep. Agrie, Agrie. Monogr. 18,

233 p.

Wolf, Frederick A., and W. J. Barbour.

1941. Brown-spot needle disease of pines. Phytopathology 31:61-74.

71
21. DOTHISTROMA NEEDLE BLIGHT OF PINES

Scirrhia pint Funk & Parker,

Dothistroma pini Hulbary

GLENN W. PETERSON

Dothistroma needle blight is the most damaging disease of

pines in the southern hemisphere (East Africa, New Zealand,

Chile). Severe damage has also been caused by Dothistroma pini

in North America, especially in plantings. Although infection

of pine seedlings in North American nurseries has seldom been

observed, it has become evident that infection in new plantings

has frequently resulted from use of infected nursery stock.

Distribution and Hosts

Dothistroma pini has been found in 23 of 50 States in the

United States and in three Provinces in Canada ; it has not been

reported in Mexico. Twenty pine species and hybrids have been

reported as hosts in North America. Dothistroma needle blight

has caused considerable consternation in the past decade because

of the devastation caused in pine plantations (chiefly Pintis

radiata) in the Southern Hemisphere.

The fungus has commonly been found on 4- to 5-year-old

pines in nurseries in the central United States which produce

pines for landscape plantings. Seedlings infected with D, pini

have seldom been detected. Yet, experience with epidemics in

new plantings in the Great Plains indicates that trees infected

in the nursery have been responsible for outbreaks in the

plantings.

Symptoms and Life Cycle

Symptoms develop in the fall of the year of infection in the

central United States and in British Columbia. Early symptoms

consist of yellow and tan spots, and bands that appear water

soaked, on the needles (fig. 21-1). The bands and spots may

turn brown to reddish brown. Commonly, the distal ends of infected needles become chlorotic, then necrotic, with the base of

the needles remaining green. Extensive necrosis of needles may

occur within 2 to 3 weeks of the first appearance of symptoms.

72
F-622525; F-522626

Figure 21-1.—Symptoms of Dothistroma needle blight on Austrian pine

needles.

Infected needles are cast prematurely. Infected second-year

needles are cast before infected first-year needles. In some seasons, second-year needles are cast extensively in late fall of the

year they became infected. In other seasons, casting is not extensive until late spring or early summer. Infected first-year

needles often are not shed until late summer follovs^ing the year

they were infected.

73
The fungus has both a sexual stage (Sdrrhia pini) and a

nonsexual stage (Dothistroma pini). The sexual stage has been

found in British Columbia, Alaska, and California, but not yet

elsewhere in North America. Ascospores are produced by the

sexual stage; their role in development of epidemics is not

known. However, much is known about the role of conidia (produced by the asexual stage) in disease development. Conidia are

cylindrical, curved, 1- to 5- but usually 3-septate, and hyaline

(fig. 21-2). The conidia are about 3.5 ^^m wide and vary in

length from 16 to 64 /^m. Conidia from western North America

are considerably longer on the average than are conidia found

elsewhere on the continent.

Conidia (spores) of D. pini are borne in stromata on. needles.

Stromata may begin to develop in the fall in the central United

States and British Columbia, but most stromata mature in the

spring following the year of infection. Stroma development in

the fall may be sufficient to cause splits in the needle epidermis,

but spores have not been observed in these stromata in the fall

(fig. 21-3). The conidia are disseminated by rainsplash throughout the growing season (May-October). Infection occurs

throughout the growing season; symptoms do not develop, however, until late summer or early fall in the central United States.

Two growing seasons are required for completion of the life

cycle in most areas of North America, although in California

the cycle may be completed in 1 year.

Control

Copper fungicides effectively control D. pini. Bordeaux mixture (8-8-100) applied twice in the growing season (mid-May,

mid-June) has provided essentially complete control in shelterbelt, Christmas tree, and other plantings in the central United

States. The first application protects previous seasons' needles;

F-522627

Figure 21-2.—Conidia (spores) of

Dothistroma pini obtained from

needles of Austrian pine.

F-522B28

Figure 21-3.—Fruiting body of Dothistroma pini raising the epidermis

of an Austrian pine needle.

74
the second application protects current-year needles. The second

application may be delayed until mid-June when controlling

this disease in plantings of Austrian or ponderosa pines, since

new needles of these species are resistant to infection until midsummer (July).

Effective control has also been obtained in plantings in the

Midwest with a single application made after considerable

growth has occurred (early June). There is some risk in this

procedure, since infection could occur prior to the early June

application. A single application will control this disease on

trees which do not have susceptible first-year needles. The above

control measures can be used by nurserymen to control Dothistroma blight in nursery windbreaks.

Detection of infected seedlings in the nursery may be difficult for two reasons: the level of infection is likely to be low,

and symptoms do not develop until late summer or early fall.

Although bordeaux mixture will effectively control this disease

in seedling beds, no tests have been made in nurseries to determine the number and timing of applications that would be required. The two-spray schedule so effective on older trees might

not be effective on seedlings. In New Zealand, monthly applications of bordeaux mixture effectively controlled the disease in

seedling beds of P. radiata.

Selected References

Cobb, F. W., Jr., and D. R. Miller.

1968. Hosts and geographic distribution of Scirrhia pini—the cause of

red band needle blight in California. J. For. 66:930-933.

Funk, A., and A. K. Parker.

1966. Scirrhia pini n. sp., the perfect state of Dothistroma pini Hulbary.

Can. J. Bot. 44:1171-1176.

Hulbary, Robert L.

1941. A needle blight of Austrian pine. 111. Nat. Hist. Surv. Bull.

21:231-236.

Jancarik, V.

1969. Control of Dothistroma pini in forest nurseries. N. Z. For. Serv.

For. Res. Inst. Res. Leafl. 24, 4 p.

Peterson, Glenn W.

1967. Dothistroma needle blight of Austrian and ponderosa pines: Epidemiology and control. Phytopathology 57:437-441.

Peterson, Glenn W.

1967. Dothistroma needle blight of pines in North America. XIV Int.

Union For. Res. Organ. Congr. [Munich, Sept. 1967] Proc, Vol. V,

p. 269-278.

Peterson, Glenn W.

1973. Infection of Austrian and ponderosa pines by Dothistroma pini

in eastern Nebraska. Phytopathology 63:1060-1063.

Thyr, B. D., and C. Gardner Shaw.

1964. Indentity of the fungus causing red band disease on pines.

Mycologia 56:103-109.

75
22. PHOMOPSIS BLIGHT OF JUNIPERS

Phomopsis juniperouora Hahn

GLENN W. PETERSON

Blight caused by the fungus Phomopsis juniperovora is a

serious nursery disease of junipers in the United States. Epidemics in nurseries in the Great Plains have resulted in total

losses in seedling and transplant beds.

Distribution

Phomopsis juniperovora is common in the Great Plains from

North Dakota to Texas, and eastward to the Atlantic coast. It

is also known to occur in Ontario.

Hosts and Damage

Members of the Cupressaceae belonging to the genera Chamaecyparis, Cupressus, Juniperus, and Thuja are infected by the

fungus. Losses have been most severe in seedling and transplant

beds of eastern redcedar and Rocky Mountain juniper (fig.

22-1). High mortality can be expected in outplantings if blighted

stock is used.

Small yellow spots appear on needles of eastern redcedar and

Rocky Mountain juniper 3 to 5 days after infection. The fungus

permeates needles and rapidly invades young stem tissue of

seedlings and transplants. As a result, terminals and branches

become light in color, then red brown, and finally ashen gray.

Lesions on stem tissue frequently develop into cankers, which

may girdle small stems and branches 1/3 inch or less in diameter. The main stem is frequently girdled at the base of infected branches. One-year-old infected seedlings are usually

killed, but older stock is less likely to be killed because the causal

fungus does not spread far below girdling cankers.

Damage due to drought may be confused with Phomopsis

blight, since in both cases tips of branches may be killed. It is

easy to distinguish the damage, however, as the line of demarcation between green and dead tissues will be sharp in Phomopsisblighted seedlings and gradual in seedlings affected by drought.

76
J.

F-B22529

Figure 22-1.—Phomopsis blight infection of second-year seedlings of eastern

redcedar.

Damage from the lesser cornstalk borer can be distinguished

from Phomopsis blight by the straw color of the dead tops and

by the feeding wounds on the lower stem and taproot. Another

fungus, Cercospora sequoiae, causes a blight of junipers and

other species in the Cupressaceae. This disease can be easily

distinguished from Phomopsis blight, since Cercospora affects

only the needles and infection starts on the oldest needles of

lower branches and spreads upward and outward, whereas

Phomopsis infection starts in newly developed needles.

Life History

The most important sources of inoculum early in the growing

season are spores produced in fruiting bodies (pycnidia) which

are formed on leaves and stems of seedlings infected the previous year. Spores are dispersed primarily by rain splash.

Young nonwounded needles are susceptible. If spores land on

these young needles, infection can occur if there is a short

period (7 hours) of 100 percent relative humidity. Spore germination, germ tube development, and infection are optimum near

24° C ; however, disease development is enhanced by higher

temperatures (32° C). Symptoms may develop within 3 to 5

days. Pycnidia with viable spores may develop within 3 to 4

weeks. Pycnidia develop not only on the infected needles, but

also on stems to which the fungus has spread from the young

needles. Pycnidia usually are not well developed until infected

77
tissues have dried considerably. They are most commonly found

on tissues which have turned ashen gray. The pycnidia are at

first embedded in needles and stems, but later partially erupt

through the epidermis. Spores are extruded in whitish tendrils.

Two types of spores (A- and B-) develop in the same or different pycnidia. A-spores are colorless, one-celled, ellipsoid, contain

two oil globules, and commonly are 7.5 to 10 by 2.2 to 2.8 /xm;

B-spores are colorless, one celled, filamentous, slightly curved,

and commonly 20.2 to 26.9 by 1 /xm. Intermediate type spores

occur infrequently. Infection is caused by the A-spores; the

B-spores do not germinate. The fungus can persist as long as

2 years as mycelium in dead parts of infected plants and produce spores. The perfect (sexual) stage of the fungus is unknown.

Control

Seedlings and transplants need protection during the entire

growing season, since spores are disseminated throughout the

period and young foliage of junipers is susceptible. Phomopsis

blight has been effectively controlled in nurseries by frequent

(usually weekly) applications of phenyl mercury fungicides.

Now that mercury fungicides cannot be used, other effective

fungicides need to be found. No effective nonmercury fungicides were found in earlier tests of over 40 fungicides. In recent

tests, benomyl have been effective under low infection conditions ; however, there is evidence that it may not be effective on

seedlings under high infection conditions. Benomyl is now registered for use in controlling Phomopsis blight.

Protective fungicides need to be applied frequently to protect

susceptible new foliage. To obtain control by limited applications, systemic fungicides would likely have to be used. Fungicide applications could possibly be limited to periods in which

flushes of new growth occur. Commonly, new growth flushes

on eastern redcedar occur early in the growing season and

again late in the summer in nurseries in the central and northern Great Plains, but during most of the growing season in the

South. Research is needed to determine if limiting applications

of protective fungicides to such periods would be effective.

Juniper seed should not be sown adjacent to beds containing

juniper stock. Poorly drained areas should also be avoided. If

overhead sprinklers are used, seedlings should be irrigated so

that water on seedlings dries before nightfall. Because shading

frames increase the length of time moisture remains on foliage,

they should not be used unless absolutely necessary. Junipers or

other hosts of this fungus should not be used in nursery wind

breaks or in landscape plantings on nursery grounds, since they

may be a source of inoculum (spores) for nursery stock. Such

trees are likely to be extensively infected if pruning results in

development of juvenile foliage.

78
Selected References

Hodges, C. S., and H. J. Green.

1961. Survival in the plantation of eastern redcedar seedlings infected

with Phomopsis blight in the nursery. Plant Dis. Rep. 45:134-136.

Peterson, Glenn W.

1965. Field survival and growth of Phomopsis-blighted and non-blighted

eastern redcedar plant stock. Plant Dis. Rep. 49:121-123.

Peterson, Glenn W.

1972. Chemical control of Phomopsis blight of junipers: a search for

new methods. Tree Planters' Notes 23(3) :3-4.

Peterson, Glenn W.

1973. Infection of Juniperus virginiana on J. scopulorum by Phomopsis

juniperovora. Phytopathology 63:246-251.

Peterson, Glenn W., D. R. Sumner, and C. Norman.

1965. Control of Phomopsis blight of eastern redcedar seedlings. Plant

Dis. Rep. 49:529-531.

Schoeneweiss, Donald R.

1969. Susceptibility of evergreen hosts to the juniper blight fungus,

Phomopsis juniperovora, under epidemic conditions. J. Am. Soc.

Hort. Sei. 94(6):609-611.

79
23. CERCOSPORA BLIGHT OF JUNIPER

Cercospora sequoiae Ell. & Ev.

CHARLES S. HODGES

Cercospora sequoiae commonly causes a needle blight in Arizona cypress in Christmas tree and ornamental plantings

throughout the southern United States, but is seldom noted on

eastern redcedar in such plantings. Occasionally, however, C.

sequoiae will cause severe damage to redcedar seedlings in the

nursery.

Distribution and Hosts

Cercospora sequoiae has been reported from most of the southern United States and as far north as Pennsylvania. Eastern

redcedar and giant sequoia are the only known seedling hosts

of the fungus. Several species of cypress, especially Arizona

cypress, Italian cypress, and Monterey cypress, and Oriental

arborvitae are susceptible in the field.

Figure 23-1.

F-522630

-Eastern redcedar seedlings infected with Cercospora sequoiae.

80
Symptoms

The disease first aiïects needles adjacent to the stem on the

lowest branches. Infection then progresses upward and outward until only the needles at the tips of the branches remain

green on severely affected plants (fig. 23-1). The disease is thus

easily distinguished from the more common Phomopsis blight,

Phomopsis juniperovora Hahn, which kills the tips of the

branches.

Life Cycle

Stromata of the fungus form profusely on the needles shortly

after they turn brown, appearing as tiny dark pustules about

100 /xm in diameter. Several brown, geniculate, 50-125 ¡xm high

conidiophores arise from each stroma (fig. 23-2). The conidia

are yellow brown, cylindrical, 5 to 6 septate, echinulate, and average about 40 X 5 ^tm. They are produced dry and are spread by

wind.

In all cases where redcedar seedlings are infected in the nursery, heavily infected Arizona cypress trees were present in the

vicinity of the beds. These infected trees probably served as

sources of infection for the seedlings. The fungus overwinters

on infected needles on living trees.

F-B22531

Figure 23-2.—Conidiophores (A) and conidia (Ä) of Cercospora seguoiae.

81
Vy^/Jll. 1/J. VFX

Seedlings should be sprayed with 8-8-100 bordeaux mixture

at 7- to 10-day intervals, once the disease is determined to be

present. Where it occurs frequently, this spray schedule can be

made standard practice, starting June 1. Sprays should be continued throughout the summer.

All Arizona cypress trees around the nursery should be examined for the disease, and infected trees removed or sprayed.

Selected References

Hodges, Charles S.

1961. New hosts for Cercospora thujina Plakidas. Plant Dis. Rep

45:745.

Hodges, Charles S.

1962. Comparison of four similar fungi from Juniperus and related

conifers. Mycologia 54; 62-69.

82
24. ROSELLINIA NEEDLE BLIGHT

Rosellinia herpotrichoides Hept. & Davis

RICHARD S. SMITH, JR.

Rosellinia needle blight was originally described as a needle

blight of mature eastern hemlock in the forests of the southeastern United States. Since the early 1950's, it has appeared

occasionally on seedlings in the forest nurseries of western

North America.

Distribution

This disease appears to be limited to California, Oregon,

Washington, and British Columbia.

WEB|r V.

|pr'>?

w

r

F-B22532

Figure 24-1.—A 2-year-oId Douglasflr seedling attacked by Rosellinia

herpotrichoides shows defoliation,

and matting of the remaining

foliage.

P-622533

Figure 24-2.—Perithecia of Rosellinia

herpotrichoides grouped on stem

and lower needles of an infected

Douglas-fir seedling.

83
Hosts and Damage

In the nurseries of western North America, the disease has

been reported on Douglas-fir and Sitka spruce.

This disease appears to kill few seedlings. The pathogen grows

over the outside of the seedling, infecting and killing needles

and small branches. The infected needles fall, and in many cases

the seedlings may be completely defoliated (fig. 24-1). This defoliation causes many seedlings to be culled, which results in the

greatest loss.

Life History

Rosellinia needle blight usually starts in the lower portions of

the seedling crown in the center of crowded beds. The fungus

grows over the outside of the seedling, infecting individual

needles as it comes to them. The fungus can be seen on the

seedling surfaces. At the advancing front of the infection, the

mycelium forms a fine white mycelial web which encompasses

whole tufts of needles and small branches. The needles then

turn yellow and die. As the mycelium ages, it and the dead

needles darken, become dense, and flatten, forming dark brown

wet mats of mycelium and needles (fig. 24-1). Groups of perithecia may be found on these old mats at the base of the seedling crown (fig. 24-2). The perithecia produce ascospores which

may be released to start new centers of infection.

This disease requires mild temperatures and long, continuous

periods of high humidity to become damaging. It usually appears after long periods of rain or fog, and becomes most severe

in the center rows of dense, overstocked nursery beds.

Control

Reduced stocking to improve aeration and to lower humidity

within the seedling beds is one of the most effective means of

controlling this disease. Some standard fungicides have been

used successfully.

Selected References

Hepting, George H., and Ross W. Davidson.

1937. A leaf and twig disease of hemlock caused by a new species of

Rosellinia. Phytopathology 27:305-310.

Shea, Keith R.

1964. Rosellinia herpotrichoides on Sitka spruce seedlings in Washington

Plant Dis. Rep. 48:512-513.

Smith, Richard S., Jr.

18966. Rosellinia needk blight of Douglas-fir in California. Plant Dis

Rep. 50:249-250.

84
25. SNOW BLIGHT OF CONIFERS

Phacidium sp., Lophophacidium sp.,

Sarcotrichila sp., Hemiphacidium sp.

DARROLL D. SKILLING

Tree seedlings are frequently damaged severely by snow blight

in the nursery. Damage is primarily limited to pines, spruces, and

firs. Snow blight in North America was formerly attributed to the

fungus Phacidium infestans, the cause of snow blight on Scotch

pine in Europe. Recent taxonomic work has shown that snow

blight in the United States and Canada is due to a number of

different fungi.

F-622534

Figure 25-1.—White spruce seedlings that show symptoms of snow blight in

early spring.

85
Distribution

Snow blight occurs in the northern United States and Canada

wherever deep snow covers the nursery beds for long periods. Snow

blight fungi are apparently native to North America, and are

widely distributed in natural stands of spruce and fir.

Hosts and Damage

The principal hosts of snow blight in North America are Norway, white, black, blue, red and Engelmann spruces ; Douglas-fir ;

ponderosa, white, and limber pines ; and balsam, white, grand, and

subalpine firs.

The fungi that collectively are called snow molds can grow at the

low temperatures found under deep snow. These include Phacidium

abietis, Lophophacidium hyperboreum, Sarcotrichila piniperda,

S. balsameae, and Hemiphacidium planum.

Foliage infected with snow blight fungi turns a glaucous-brown

color in the spring after the snow melts (fig. 25-1). Infection

frequently occurs in sharply defined patches, and is usually most

severe in nursery areas where snowmelt is delayed. After the

snow melts, a covering of white mycelium can be seen on the

browning foliage. Mild attacks kill only the foliage below the

snowline, but heavy infections kill large numbers of seedlings. Frequently, however, the roots live for some time after the shoots

are dead.

Life History

In late summer, small, brownish-black, disk-shaped apothecia

appear on the undersurface of infected, browned needles. Ascospores are discharged from these apothecia during moist weather

until winter. Tiny, black microsclerotia, resistant to drought and

other unfavorable climatic factors, are also produced on the surface

of infected needles.

Primary infection of needles occurs in the fall from the winddisseminated ascospores. The ascospores resting on the needle

surfaces germinate as soon as the needles become embedded in

snow, from which the spores receive the required moisture. Needles

of all ages are susceptible. The microsclerotia probably also

function like spores. Because the fungus is able to grow under

a deep snow cover, seedlings that appear healthy in the fall may

show severe foliage browning or mortality the following spring.

Secondary infections also occur, as the fungal mycelium from

infected needles grows out under the snow onto adjacent foliage

and enters healthy needles.

Control

Snow blights may occur inconspicuously on low branches of

trees up to the pole size; in spruces, even later. Therefore, care

86
should be taken to minimize such sources of infection near or

within a nursery.

As far as possible, localities where snowmelt is delayed should

not be used as nursery sites. Moreover, susceptible tree species

should not be grown in nursery beds where snow tends to accumulate in drifts. If only scattered patches of this disease are found

in nurseries, its spread may be eliminated by careful removal of

visibly infected seedlings and those adjacent to them.

In Sweden, seedlings from a northern provenance are more

resistant to snow blight than those from a southern provenance.

Finnish workers found that snow blight is severe in nurseries

where the soil is low in potassium.

Selected References

Baxter, Dow Vawter.

1952. Pathology in forest practice. 2d Ed., 601 p. John Wiley & Sons,

N.Y.

Björkman, Erik.

1963. Snow blight of pines, p. 70-72. In Internationally dangerous forest

tree diseases. U.S. Dep. Agrie. Misc. Publ. 939, 122 p.

Boyce, John Shaw.

1961. Forest pathology. 3d Ed., 572 p. McGraw-Hill, N.Y.

Reid, James, and R. F. Cain.

1962. Studies on the organisms associated with "snow-blight" of conifers

in North America. II. Some species of the genera Phacidium,

Lophophacidium, Sarcotrichila, and Hemiphacidiwm. Mycologia 54:

481-497.

87
26. SYCAMORE ANTHRACNOSE

Gnomonia platani Kleb.,

Gloeosporium platani Oud.

WiNAND K. HOCK AND FREDERICK H. BERRY*

Sycamore anthracnose, caused by the fungus Gnomonia platani

Kleb, (asexual stage, Gloeosporium platani Oud.), is often responsible for the shabby and ragged appearance of sycamores

during May and June. Although usually more unsightly than lethal,

repeated defoliation by this disease for several successive years

can so weaken trees that they may succumb to other pests or to

environmental pollutants and climatic stress.

Distribution

Anthracnose occurs throughout the entire range of sycamores.

Hosts and Damage

The disease is extremely severe in some years on American and

California sycamores, although all species of sycamore—including

the less susceptible London plane tree—can be attacked when conditions are suitable for the fungus.

Four distinct stages of sycamore anthracnose are generally

recognized: twig, bud, shoot, and leaf blight. Any one or more of

these stages can be seen during the growing season, although most

damage to nursery seedlings results from the leaf blight stage

unless plants are grown for more than one season in the nursery.

The entire developing shoot of nursery seedlings may die. This

shoot blight is probably the result of direct infection and is

comparable to the leaf blight stage of older trees.

The twig and bud blight stages develop early in the spring while

the trees are still dormant. Cankers develop on infected twigs and

ultimately girdle and kill the buds and tips of the twigs. Often

the buds are killed even before they begin to swell. Repeated

killing of twigs throughout the crown results in a gnarled and

abnormally branched tree. Small, black fruiting bodies (acervuli)

of the fungus appear in the bark of infected twigs and branch

cankers.

* Plant Pathologist, Pest Management Section, Division of Plant Industries, Maryland Department of Agriculture, College Park, Md., and

Principal Plant Pathologist, Northeastern Forest Experiment Station, USDA

Forest Service, Delaware, Ohio.

88
Shoot blight appears as the sudden dying of developing shoots

and immature leaves. The fungus girdles and kills tissues surrounding the shoots, which kills the young leaves on the emerging

shoots. This damage is often confused with frost injury.

The leaf blight stage is produced by direct infection of the

developing or mature leaves by fungus spores produced on twigs

and branch cankers. Irregular, necrotic areas form adjacent to

and around diseased spots on the midrib and principal veins (fig.

26-1). Creamy brown pustules of the fungus are evident on the

diseased tissues, particularly on the midrib and veins.

. oc -I A 1. F-622536 figure 26-1.—Anthracnose of sycamore leaves causes irregular, necrotic areas

adjacent to and surrounding diseased spots on the midrib and principal veins.

(Photo courtesy of Illinois Natural History Survey, Urbana, Illinois).

The disease is severe in some years and virtually nonexistent in

others. Temperature is critical during the first 2 weeks after leaf

emergence. Shoot blight is usually severe if mean daily temperatures average between 10° and 13° C during this period. Shoot

bhght decreases as the average temperature increases above 13° C.

Life History

The principal sources of inoculum for early spring infections

are conidia produced on infected twigs and branches and on fallen

leaves. Ascospores borne in perithecia are also produced on fallen

leaves under certain conditions. Conidia and ascospores are carried

from infected twigs and leaves by wind and splashing rain to

new leaves, where they germinate and cause new infections. Conidia, borne in structures called acervuli, are hyaline, one celled.

89
ovoid to oblong, sometimes slightly curved, and average 9.7 x 5.0

fxm in size. Ascospores, on the other hand, are asymmetric in shape,

have rounded ends and two cells, the smaller cell being only about

F-B22537; F-522538

Figure 26-2.—Above, Control of sycamore anthracnose on 1-year-old American

sycamore seedlings with two applications of triarimol fungicide; Below,

unsprayed control plot.

90
one-fifth to one-fourth the length of the larger. Ascospores average

13.4 X 3.7 /xm in size.

During wet weather, small creamy brown pustules containing

conidia develop in the underside of the leaf on dead tissue along

the veins. These conidia may be washed to other leaves by rain

to cause new infections. The fungus then spreads through the

veins, midribs, and petioles into the twigs, where it overwinters

and becomes active the following spring. The cycle is repeated when

acervuli appear in the bark of infected twigs, and conidia are

released to initiate new infections.

Control

There are two control procedures: sanitation and chemical

sprays.

Fallen leaves and twigs should be gathered and burned before

the fungus becomes active in the spring. Infected twigs and

branches should be pruned and burned whenever possible. Application of a complete fertilizer will improve the vigor of trees

weakened by repeated attacks.

Control of sycamore anthracnose is possible with properly timed

applications of a suitable fungicide. Current recommendations call

for three or more applications of zineb, dodine, or copper fungicide.

The first spray should coincide with a swelling of the buds, and

the second and third should be made at approximately 10- to

14-day intervals after the initial spray.

Newer experimental fungicides such as triarimol, benomyl, and

thiophanate-methyl have shown considerable promise for combating sycamore anthracnose in an Ohio nursery (fig. 26-2). They

may eventually replace the current generation of materials, which

are less effective than the environmentally unacceptable organic

mercury compounds.

Selected References

1964.' Illinois trees: Their diseases. 111. Nat. Hist. Surv. Circ. 46, 96 p.

1971. Diseases of forest and shade trees of the United States. U.S. Dep.

Agrie, Agrie. Handb. 386, 658 p.

Hock, Winand K. . ., x x 14-4? 1971. Sycamore anthracnose. Fungicide and nematicide tests, results oí

1971. 27:138-139. Am. Phytopath. Soc, Washington, D.C.

Neely, Dan, and E. B. Himelick. «, 4. n- u

1963. Temperature and sycamore anthracnose severity. Plant Dis. Kep

47:171-175.

Pirone, Pascal Pompey.

1972. Tree maintenance. 574 p. Oxford Univ. Press, N.Y.

Schuldt, Paul H. , . ^,

1955. Comparison of anthracnose fungi on oak, sycamore, and other

trees. Contrib. Boyce Thompson Inst. 18(2) :85-107.

Sinclair, W. A., and W. T. Johnson. .T .r ox x /^ ,1 A •

1968. Anthracnose diseases of trees and shrubs. N. Y. State Coll. Agric,

Cornell Tree Pest Leafl. A-2, 7 p.

91
27. WALNUT ANTHRACNOSE

Gnomonia leptostyla (Fr.) Ces. & de N.

FREDERICK H. BERRY AND WINAND K. HOCK

Walnut anthracnose, sometimes called leaf blotch, is a destructive disease of certain walnut species, particularly the eastern black

walnut. It IS caused by the fungus Gnomonia leptostyla (Fr.)

ÏT'i "^^ /xî'xîf imperfect stage of which is Marssomna

mglandis (Lib.) Magn. The disease may quickly reach epidemic

proportions during wet weather in the growing season and

cause many walnut trees-from seedlings to mature trees-to

lose almost all their leaves by late July or early August This

premature defoliation slows growth, weakens trees, and sometimes

kills them Nuts from infected trees are often poorly filled and

01 low quality.

Distribution

Walnut anthracnose is distributed worldwide. Besides infecting Juglans species in the United States, the disease has been

reported from most of the countries of Europe and from Argentina, Canada, and South Africa. It is likely to be present in all

nurseries where black walnut is grown.

Hosts and Damage

Eastern black walnut trees vary in their susceptibility to walnut

anthracnose. But when weather favors development of the fungus

Tr". 5^TJ^!? susceptible trees become severely infected and defoliated. Butternut, Persian (English) walnut, and first-generation

hybrids of Persian and black walnuts are also attacked. Two species

of walnut native to California, the Hinds walnut and the California walnut, are both susceptible.

Leaves, nuts, and occasionally shoots of the current season's

growth are attacked by the fungus. Tiny dark brown or black

%° o^'^'i^" -iu*'' irregularly circular, appear on infected leaves

(ng. ¿1-1). These spots gradually become more numerous, enlarge, and often merge to form still larger dead areas The leaf

area bordering these spots is usually chlorotic. Most infected

leaves and leaflets fall prematurely, but a few sometimes remain

attached to the tree for most of the season.

92
F-522B39

Figure 27-1.—Anthracnose lesions on a black walnut leaf.

Sunken, necrotic spots, usually smaller than those on the leaves,

appear on the husks of infected nuts. When immature nuts become diseased, they do not develop normally, and may drop

prematurely.

Lesions are occasionally found on current shoots. These lesions

are dead, sunken areas, oval to irregularly circular.

Life History

The fungus usually overwinters in fallen walnut leaves infected

during the preceding summer. The fungus develops in these diseased leaves, and produces ascospores in the spring. These ascospores, which at maturity are 2.5 to 4.0 by 18.0 to 25.0 ¡xm in

size, cause primary infection. They are discharged from the walnut leaves during rainy periods and carried upward by wind or a

combination of wind and rain. If they lodge on a susceptible

leaf under favorable conditions, the ascospores germinate; leaf

spots appear in about 14 to 16 days.

Secondary spores, the conidia, are produced on diseased leaflets

in black fruiting bodies called acervuli. Acervuli are more abundant on the under side of leaflets, but occasionally a few are found

on the upper side. The conidia are colorless, usually crescent

shaped, and divided by the crosswall into two approximately equal

cells. They are 3 to 4 by 14 to 30 ^^m in size. Conidia are borne

in large numbers, and are spread from leaf to leaf by wind and

splashing rain. The rapid increase and spread of walnut anthracnose in the summer and fall is attributed to repeated generations of conidia. Leaves are most likely to be infected and fall

off during wet weather.

93
Control

Walnut anthracnose can be controlled in forest nurseries by

periodic spraying with fungicides. Dodine (Cyprex) effectively

controlled the disease. To protect the seedling trees from primary

infection, the first spray must be applied before ascospores are

discharged. The foliage on black walnuts is about one-half mature

size at this time. Then, depending on weather conditions, two or

three additional sprays should be applied at approximately 2-week

intervals.

Recently an experimental fungicide, triarimol, has shown promise for controlling walnut anthracnose in Ohio nursery beds (fig.

27-2). Further studies are in progress to test its effectiveness.

F-622540

Figure 27-2.—Control of anthracnose on black walnut seedlings with triarimol

fungicide. Unsprayed plots in foreground.

Selected References

Berry, Frederick H.

1960. Etiology and control of walnut anthracnose. Md. Agrie. Exp. Stn.

Bull. A-113, 22 p.

Berry, Frederick H.

1970. Walnut anthracnose. U. S. Dep. Agrie. For. Pest Leafl. 85, 4 p.

94
28. MARSSONINA BLIGHT OF BIGTOOTH

ASPEN

Marssonina populi (Lib.) Magn.

LEON S. DOCHINGER*

A foliar blight caused by the fungus Marssonina populi

(Lib.) Magn. is the most damaging disease of bigtooth aspen

seedhngs in Ohio forest nurseries. Occasionally plant mortality

exceeds 90 percent in 1-year-old beds. Also, weakened trees do

not survive when outplanted, and may serve as a source of inoculum

for infecting mature poplars and aspens.

Distribution

Although Marssonina leaf blight of bigtooth aspen seedlings is

reported only in Ohio forest nurseries, the organism is common on

F-522541

Figure 28-1.—Marssonina leaf blight

symptoms on bigtooth aspen.

F-B22642

Figure 28-2.—Marssonina blighted

bigtooth aspen seedling.

* Principal Plant Pathologist, Northeastern Forest Experiment Station,

USDA Forest Service, Delaware, Ohio.

95
trees throughout Canada and the United States. The causal fungus

attacks aspen and poplar species in California, the southern Rocky

Mountain region, the Intermountain region (particularly Utah,

western Wyoming, and southeastern Idaho), the North Central

region (Iowa and Indiana), most of the Northeastern region, and

into southern Canada.

Hosts and Damage

Criteria are insufficient for separating the various host-specific

strains of Marssonina into more than a single species. Thus, all

reported pathogenic species of Marssonina on poplar hosts will be

considered here as M. populL

In North America, M. populi is associated with leaf diseases on

practically all native and exotic aspen and poplar species. Hosts of

this organism include white poplar, bigtooth aspen, and quaking

aspen of the genus Populus in section Leuce ; eastern cottonwood

and black poplar in section Aigieros ; and Carolina poplar, balsam

poplar, narrowleaf cottonwood, and black cottonwood in section

Tacamahaca.

M. populi produces two types of leaf spots on bigtooth aspen

seedlings. On more tolerant plants, superficial brownish spots

about 1 mm in diameter develop over the entire surface of the

leaf and at times on the petiole. On susceptible hosts, these minute

spots occur along with irregular reddish-brown lesions (4 to 5 mm

in diameter) which penetrate both sides of the leaf (fig. 28-1).

The infection process is accelerated by repeated rainstorms and

increasing temperatures during the late spring and early summer

months. With such favorable environmental conditions for disease

development, lesions coalesce into rust-brown to blackened necrotic

areas in which the fungus sporulates abundantly. Lower leaves

absciss prematurely and defoliation progresses upwards toward

the tips of the shoots. When infection becomes severe, shoots are

blighted and seedling mortality results (fig. 28-2, 28-3).

Life History

The leaf blight organism overwinters on dead twigs and plant

refuse. By mid-May, conidia from hibernating stroma provide

primary sources of inoculum. Airborne conidia are probably

responsible for most of the initial leaf infections of bigtooth aspen

seedlings. In nursery beds disease development is conditioned by

climatic factors. Maximum damage may occur within a temperature range of 15°-25° C and 30-44 mm rainfall.

Later in the season, acervuli are produced on the lesions of

diseased leaves. From these fruiting pustules, both macroconidia

and microconidia are exuded in whitish-yellow masses. Conidia

are disseminated from the action of rain splash, and cause infections of bigtooth aspen plants throughout the nursery beds.

96
r-B22543

Figure 28-3.—Marssonina blight can devastate entire beds of bigtooth aspen.

Control

Cultural practices are important in reducing losses attributable

to Marssonina leaf blight. These control practices include (1) removal of infectious plant debris, (2) reducing the planting density

of seedlings for better air movement, and (3) avoiding overhead

irrigation systems.

F-522544

Figure 28-4.—Bigtooth aspen seedlings protected by fungicide (Benlate).

97
Recently, Benlate has shown promise of controlling infections

on bigtooth aspen in Ohio nursery beds (fig. 28-4). Studies of its

effectiveness are continuing for possible registration.

Since some seedlings tolerate M. populi, natural selection and

breeding of trees should be examined. Resistant clonal material

under varying environments should also be investigated.

Selected References

Boyer, M. G.

1961. Variability and hypal anastomoses in host-specific forms of

Marssonina populi (Lib.) Magn. Can. J. Bot. 39:1409-1427.

Castellani, E.

1966. Inñuence of climatic conditions on infection of M. brunnea of

Euro-American poplars. Phytopathol. Mediterr. 5:41-52.

Gremmen, J.

1965. The Marssonina disease of poplar. 3. The occurrence of M. brunnea

in the Netherlands. Ned. Bosb.-Tijdscher 37:196-198.

Magnani, G.

1964. Poplar leaf alterations by Marsonninae species. Pubbl. Cent. Spec.

Agrie. For. 7:251-283.

Mielke, James L.

1957. Aspen leaf blight in the intermountain region. U.S. Dep. Agrie.

For. Serv. Intermt. For. Exp. Stn. Res. Note 42, 5 p.

98
29. MELAMPSORA RUST ON COTTONWOOD

Melampsora medusae Thum.

T. H. FILER, JR.

Melampsora rust has been known to afïïict native and introduced poplars in the northern United States since the turn of the

century, but its presence in the lower Mississippi River valley

was not recognized until 1967. Each year thousands of acres in

this area are being planted to cottonwood, and leaf rust infections

in these plantations cause defoliation prior to normal abscission

in the fall. In some years defoliation occurs in summer, reducing

growth and vigor of planting stock.

Distribution

Melampsora rust is widespread in Canada and the United

States. Severe summer defoliation of nursery stock has been reported in Mississippi, Illinois, and Texas.

Hosts and Damage

Eastern cottonwood, quaking aspen, balsam poplar, plains

cottonwood, Rio Grande poplar, and many hybrids are hosts of

Melampsora medusae.

The first symptoms, which may appear as early as April in

Texas, are minute chlorotic spots on upper or lower leaf surfaces.

Figure 29-1.—Melampsora medusae uredia on cottonwood leaf.

99

F-522545
The spots enlarge, and yellow, powdery pustules develop to produce urediospores (fig. 29-1), which start new infections. If

humidity is high, infections may recur until the entire leaf is

covered with yellow spores. Defoliation will result when rust

covers 50 percent or more of the leaf tissue. In a cutting nursery,

defoliation in midsummer reduces vigor and limits height growth,

thereby reducing the number of merchantable cuttings.

Life History

Melampsora requires two hosts to complete its life cycle. Cottonwood leaves are infected by windborne urediospores in spring,

summer, and fall. These spores form yellow, powdery pustules,

which release spores to cause additional cottonwood leaf infection.

In late summer, telia are formed in orange-yellow crusts in the

same pustules which formerly produced urediospores. The telia,

which overwinter on fallen leaves, change in color from yellow to

brown, and finally to black. In the spring, the germinating teliospores produce basidiospores which infect larch, Douglas-fir, and

lodgepole, ponderosa, Monterey, and Scotch pines. Teliospores

cannot infect cottonwood. Spring infection of cottonwood is by

windborne aeciospores produced on conifers, or by urediospores

that overwinter on green poplar leaves in frost-free areas.

During the summer, temperatures above 35°C and dry weather

usually limit the spread of rust. In the fall, new rust infections

occur when the micro-climates favoring rust development reappear. It appears the rust spread in fall is from the northern range

to the southern range by successive urediospore infections.

Control

Melampsora rust has been controlled in experimental nursery

tests by one summer application of cupric oxide or benomyl. A

number of rust-resistant clones developed by the Southern Hardwoods Laboratory are available through several State forest nurseries. Stoneville clones 75 and 92 are not defoliated by rust, and

should be favored in areas where summer defoliation is known

to occur.

Selected References

Chitzanidis, A., and E. P. Van Arsdel.

1970. Autumn introduction and winter survival of poplar rust on the

Texas coastal plain. (Abstr.) Phytopathology 60:582.

Filer, T. H.

1973. Hardwood nursery diseases. Forest Tree Nurserymen's Conf.

Proc. 1972:25-27. West. Session, USDA Forest Serv. SE-S&PF.

Toole, E. Richard.

1967. Melampsora medusae causes cottonwood rust in Lower Mississippi

Valley. Phytopathology 57:1361-1362.

Ziller, Wolf G.

1965. Studies of western tree rusts. VI. The aecial host ranges of

Melampsora albertensiSy M, mediisae, and M, occidentalism Can. J.

Bot. 43:217-230.

100
30. SEPTORIA LEAF SPOT AND CANKER ON

COTTONWOOD

Septoria musiva Peck

T. H. FILER, JR.

Waterman reported in 1946 that Septoria musiva Peck caused

cankers and leaf spots on hybrid poplars in the United States, but

only recently was the fungus recognized as a cause of canker

disease in cottonwood (Populiis deltoides Bartr.) (fig. 30-1).

S. musiva is currently causing high cull rates in nurseries producing cottonwood cuttings in the lower Mississippi River valley

(Mississippi, Louisiana, and Arkansas). While the canker produced by the fungus does not girdle the stem, it provides entry

courts for other fungi, such as species of Cytospora, Phomopsis,

and Fu^arium., which do girdle.

r-522547; F-522649

Figure 30-1.—Left, typical canker caused by Septoria musiva on 1-year-old

cottonwood stem. Right, leaf spots on cottonwood caused by Septoria

musiva.

101
Distribution

S. musiva occurs on poplars throughout the United States,

Canada, and Argentina. It causes cankers on cottonwood in the

lower Mississippi River valley.

Hosts and Damage

Septoria musiva causes leaf spot on ail native species of aspen

and poplars (fig. 30-1). It produces cankers on P. deltoides and P.

jackii Sarg., a natural hybrid. It also causes cankers on many

introduced species and hybrids. Since all native species of aspen

and other poplars are susceptible to leaf infection, no poplar can

be considered immune to S. musiva stem canker.

Leaf infections usually precede infection of stems. Tender

leaves are infected at bud break, and leaf spots develop 1 to 2

weeks later. Spots first appear as depressed black flecks. Under

favorable moisture conditions, flecks increase to a size of 2 to

5 mm. Spots coalesce on leaves with multiple infections, and as

much as 50 percent of the leaf tissue can be affected. As the

necrotic tissue dries, it fades to light tan or white in the center.

Since the margin remains black, the leaf spot has a ''bull's eye'*

appearance. Three to four weeks after infection, pycnidia appear

as small, black, inconspicuous flecks in the centers of leaf spots.

Spores, which exude during periods of high humidity, can infect

leaves or stems.

In new cottonwood nurseries established from unrooted cuttings,

leaf infections may occur during the first season from spores

lodged on cuttings taken from infected stool beds, and from windborne spores from nearby plantations on native poplars.

Usually, less than 1 percent of 1-year-old cottonwood nursery

stock has canker infections. Cankers are more frequent on 2- to

6-year-old stool beds. The increase in infection is due to the

ascogenous stage, which overwinters on leaves and twigs on the

ground. Cankers, which usually are within 3 feet of the ground,

first appear as slightly sunken areas, smooth in the center, with

raised margins. The infected cambium becomes black, the center

bleaches to an ashy white, and tiny black pycnidia can be observed.

The lesions average 1 cm wide by 4 cm long ; they may persist to

form perennial cankers. Stems infected with S. musiva are

rapidly attacked by other fungi that cause additional damage.

Planting stock containing cankers must be culled.

Life History

Most early spring infections on basal branches and leaves

originate from ascospores. These spores are produced in tiny,

black, flask-shaped fruiting bodies or perithecia which develop on

fallen leaves or stems. When humidity is high, a pore is formed

in the perithecia and ascospores are forcibly discharged into the

102
aiL, j.iiiuugiiuuL tiit; gruwiiig seasün, iurtner inieciion occurs

during periods of high humidity, when pycnidiospores exude from

leaves and stems. Spores are washed by rain or moved by air to

new infection sites on stems and leaves.

Control

Loss of cuttings due to cankers can be greatly reduced through

sanitation. After cuttings are harvested from stool beds in the

winter, all leaves and stems should be plowed under or removed

to prevent new shoots from being infected in spring by ascospores.

Native poplars in or near the nursery should be removed to prevent early leaf infections by airborne pycnidiospores.

Selected References

Bier, J. E.

1939. Septoria canker of introduced and native hybrid poplars. Can. J.

Res. 17 (Sect. C) : 195-204.

Filer, T. H., F. I. McCracken, C. A. Mohn, and W. K. Randall.

1971. Septoria canker on nursery stock of Populus deltoides. Plant Dis.

Rep. 55:460-463.

Sarasola, A. A.

1944. Dos septoriosis de las alamedas Argentinas. Rev. Argent, de

Agron. 11:20-43.

Waterman, Alma M.

1946. Canker of hybrid poplar clones in the United States, caused by

Septoria musiva. Phytopathology 36:148-156.

103
31. YELLOWS, OR CHLOROSIS

S. J. ROWAN

"Yellows" and "chlorosis" are terms used to describe any

physiological disturbance or disease of plants resulting in the

yellowing or blanching of foliage (fig. 31-1). The destruction or

reduced synthesis of chlorophyll may be caused by several biotic

or abiotic factors, and is symptomatic of the decline in health of a

plant. In general, chlorotic seedlings are unusually sensitive to

injury by high or low temperatures, nematodes, fungi, bacteria,

pesticides, and other factors. The interactions of chlorosis and

these secondary agents cause the annual loss of up to 50 percent of

seedling crops in forest tree nurseries. These losses result from

both seedling mortality and the culling of seedlings which do not

reach plantable size.

F-522550

Figure 31-1.—Chlorosis of loblolly (left bed) and slash (right bed) pine

seedlings.

104
uescriptíon of Damage

All green plants are subject to chlorosis. The amount of growth

reduction or mortality of seedlings depends upon the cause of the

chlorosis, the length of time the plants are chlorotic, and the

amount of chlorophyll that is deficient. Although chlorosis may be

associated with rapid necrosis and death of the root system and

plant, severe chlorosis may cause only slight growth reduction if it

IS of short duration, or it may cause high mortality if it persists

for several months.

Causes

Although two or more agents often combine to cause chlorosis

of forest tree seedlings, the more common agents are as follows:

Deficiencies or excesses of mineral elements essential to plants.

The elements most commonly associated with chlorosis in forest

tree seedlings are iron, nitrogen, phosphorus, potassium, and

calcium. Magnesium, manganese, zinc, boron, copper, molybdenum,

and sulfur may also be associated with chlorosis.

Toxic levels of chemical compounds on foliage or in soils.

Excess herbicide, fungicide, nematicide, insecticide, or other

compounds used in nurseries may induce chlorosis.

Insects and mites.—Feeding is more commonly associated with

chlorosis than is oviposition, stinging, or other activities. Red

spider mites, for example, commonly cause chlorosis of coniferous

seedlings by feeding on foliage.

Parasitic fungi, bacteria, and nematodes.—Pathogens that

cause root disease are most commonly associated with chlorotic

seedlings, but foliar and stem pathogens may also cause chlorosis.

Viruses and mycoplasms.—"Yellows" is the descriptive term

applied to those plant diseases caused by certain viruses and

mycoplasms.

Genetic abnormalities.—The lack or loss of the ability to synthesize chlorophyll in plants may be caused by mutation or may

be simply inherited. Albinism, a homozygous recessive condition,

is the most common genetic abnormality associated with chlorotic

forest tree seedlings.

Environmental stresses.—Excess soil water and high or low soil

and air temperatures may cause chlorosis. Cold is often associated

with the synthesis of pigments other than chlorophyll, leading to

red, purple, yellow, or other pigmentation.

Mycorrhizae.—A deficiency of mycorrhizae will cause chlorosis.

Excessive use of soil fumigants may reduce the number of mycorrhizal roots and cause chlorosis.

Miscellaneous agents.—Parasitic seed plants, such as dodder or

seymeria, may cause chlorosis of tree seedlings.

105
uoniroi

Because chlorosis of forest tree seedlings may be caused by any

of several factors, cause (s) need to be accurately and rapidly

diagnosed. The usual diagnostic procedure involves a process of

elimination, with obvious and easily detected causes checked first.

Excess soil water, insects, heat or cold injury, and disease symptoms are frequently easy to detect. Foliar and soil analysis will

pinpoint any major nutritional inadequacies. These nutrient deficiencies can usually be corrected by applying the requisite elementís) to the foliage or soil.

The removal of excess fertilizer or toxic chemicals from soil is

often difficult, and may require fallowing the contaminated area

until the material has deteriorated or leached below the plant root

zone by rain or irrigation. Contaminated soil may have to be

removed. Insects and mites may be controlled with insecticides.

Root diseases may be controlled by soil fumigation, before planting or after harvesting, with methyl bromide and other soil fumigants. Foliar and stem pathogens may be controlled by application

of fungicides. Viral, mycoplasmic, and certain bacterial diseases

may be controlled by excluding symptom-bearing plants from the

nursery. Mulching, shading, and irrigating seedbeds to reduce air

and soil temperatures provide some control for chlorosis caused by

high temperatures. Proper soil drainage will eliminate excess soil

water and the associated seedling chlorosis.

Selected References

Davis, William C, Ernest Wright, and Carl Hartley.

1942. Diseases of forest-tree nursery stock. U.S. Dep. Agrie, Civ.

Conserv. Corps, For. Publ. 9, 79 p.

Foster, A. A.

1959. Nursery diseases of southern pines. U.S. Dep. Agrie. For. Pest

Leafl. 32, 7 p.

Hacskaylo, J., P. F. Finn, and J. P. Vimmerstedt.

1969. Deficiency symptoms of some forest trees. Ohio Agrie. Res. and

Develop. Center Res. Bull. 1015, 69 p.

Hodges, Charles S., Jr.

1962. Black root rot of pine seedlings. Phytopathology 52:210-219.

Hodges, Charles S., Jr.

1962. Diseases of southeastern forest nurseries and their control. U.S.

Dep. Agrie. For. Serv. Southeast. For. Exp. Stn., Stn. Pap. 142,

16 p. Asheville, N.C.

Hodges, Charles S., Jr., and John L. Ruehle.

1969. Nursery diseases of southern pines. U.S. Dep. Agrie. For. Pest

Leafl. 32, 8 p.

106
32. AIR POLLUTION

LEON S. DOCHINGER AND KEITH F. JENSEN*

In the early 1900's, occasional damage to nursery crops in Canada and parts of the United States was attributed to fumes from

local industrial operations. Today, besides such acute damage, the

more insidious effects of long-term chronic pollution are being reported. Collectively, both the acute and chronic forms of air

pollution are serious threats to the welfare of forest nurseries. In

time, the primary offenders—ozone, fluorides, and sulfur dioxide—

will probably become more widespread and may cause greater

damage to forest tree nurseries. Also, new chemical forms of

pollution arising from advancing technology may combine with

current minor pollutants to cause more phytotoxicity to nursery

plants.

* Respectively, Principal Plant Pathologist, Northeastern Forest Experiment Station, USDA Forest Service, Delaware, Ohio, and Plant Physiologist,

Northeastern Forest Experiment Station, USDA Forest Service, Delaware,

Ohio.

F-5225B2

Figure 32-1.—White pine: needle burn from oxidants under field conditions.

107
Ozone

Ozone (O3) is the prime constituent of the photochemical smog

that forms near urban areas. This oxidant damages more vegetation than any other pollutant in the temperate zone of North

America.

Sources

A major portion of ozone is formed through light-energized

reactions with the emissions from industries and automobile engines. Also, ozone is transferred to the earth's surface from the

upper atmosphere by turbulence. Ozone also can be produced by

electrical storms.

Symptoms

Conifers,—Rapid intake of toxic levels of ozone causes acute

responses in forest tree species. Characteristic acute symptoms

include needle banding and tip necrosis (fig. 32-1). Earliest

manifestations from high concentrations of ozone are cellular

collapse and the subsequent appearance of necrotic tissue on all

ages of needles. Minute yellow, pink, or silver spots, along with

mottled green and yellow patterns of chloronemia, are indications

of chronic exposure to ozone. Intensification causes needle tipburn,

defoliation, dwarfing of retained needles, and reduced seedling

growth. Chronic needle symptoms result from low levels of oxidant concentrations for extended periods, and also may be observed on conifer needles displaying acute responses.

Hardwoods.—Both acute and chronic symptoms may appear on

leaves of the same seedling. High concentrations of ozone cause

necrosis on upper and lower leaf surfaces, and leaf drop of some

broad-leaved species (fig. 32-2). Chronic exposure produces

chlorotic and pigmented stipules and flecks on the upper surfaces

of the foliage. When these metallic-like lesions coalesce, they impart an overall bronzed, silvery, or purple cast to damaged foliage.

Little injury is expressed initially on some trees, but a few days

after chronic exposure to ozone, premature loss of some mottled

foliage results.

Fluorides

The fluorides are more harmful to vegetation than similar quantities of most other aerial phytotoxicants. Of the fluorides released into the atmosphere, the gaseous compounds of hydrogen

fluorides and silicon tetrafluorides are more injurious than the

particulate fluorides. Because of the expansion of industries that

emit fluorides, there is potentially greater danger to seedlings

from these pollutants if industrial control measures are ineifective.

108
^^

-*í

F-5225B3; r-B22554

Figure 32-2.—Above, Hybrid poplar: foliar injury from acute concentrations

of O3 fumigation; Below, bigtooth aspen: purple stippling and overall

bronze cast from chronic levels of O3 fumigation.

Sources

Fluorides are emitted into the atmosphere from electrolytic

reduction of alumina; combustion of coal; manufacture of brick,

phosphate fertilizer, and pottery; ferro-enamel processes; and

steel and nonferrous ore smelting.

109
Symptoms

Conifers,—Symptoms vary widely in forest seedlings injured by

gaseous fluorides. Usually, injury results from an accumulation of

fluorides in plant tissues over an extended time. Chronic symptoms

are chlorosis, banding, needle drop, and growth suppression. With

continued absorption of low level (chronic) concentrations of

fluoride, conifer needles, at first chlorotic, now become buff to

reddish brown, and needle burn and tip necrosis occur over the

entire conifer seedling. Acute damage is often characterized by

tip and needle browning. Generally, older needles are more tolerant to fluoride pollutants, whereas new foliage is susceptible to

both acute and chronic fluoride exposures.

Hardwoods,—In deciduous seedlings, fluorides are absorbed into

the leaves and accumulate along the margins and apex. Usually, an

initial chlorosis of the leaf tip develops and later extends downward along the margin and inward toward the midrib. With

continued intake of fluorides, chronic areas become necrotic, and

tissue degenerates. On some broad-leaved species, a given quantity of fluorides may cause necrosis of the more sensitive trees,

growth reduction of less susceptible trees, and growth stimulation

in more tolerant trees over similar exposure periods.

Sulfur Dioxide

For many years emissions of sulfur dioxide (SO2) have damaged forest trees. Many of these harmful effects are produced not

only by SO2 but also from related oxides of sulfur, their acids, and

acid salts. These sulfur-containing compounds may combine with

other pollutants and create more plant injury than when they

operate independently.

Sources

Primary sources of SO2 are the combustion of fossil fuels, smelting of ores, production and manufacturing of natural gas, utiHzation of sulfur-containing compounds, and refining of petroleum

products.

Symptoms

Conifers.—Sulfur dioxide may cause acute and chronic injury

to conifer seedlings (fig. 32-3). The former results from a rapid

absorption of high concentrations over a relatively short period.

All ages of foliage are affected, but not all needles in a fascicle are

harmed. Injured portions of the needle display a tannish-brown

discoloration and tip necrosis. Chronic responses are usually found

on the current year's needles. Chlorosis, necrosis, reduction in

needle length, and needle loss often occur after exposure to sublethal concentrations of SO.. Needles may also display both acute

and chronic symptoms.

110
F-522B56; F-5225B6

Figure 32-3.—Above, White pine: needle necrosis from acute levels of SO2

fumigation; Below, Scotch pine: needle mottle from chronic levels of SO2

1

in field.

Hardwoods.—After acute exposures to SO2, tissues are dull and

have a water-soaked appearance. In time, the tissues blanch and

assume a gray-green color. These affected areas dry and bleach

to a light-tan or ivory color in some species, but most often they

become reddish brown. The final pattern consists of interveinal

necrosis, but little efl'ect is discernible across the veins of leaves

unless injury is very severe. Chronic injury leads to a change in

111
leaf pigmentation, ranging from flecks to a diffuse mottling. Colors

range from yellow, ivory, and bronze to black (figs. 32-4, 32-5).

F-B22557; F-522558

Figure 32-4.—Yellow poplar, with chronic foliar symptoms of yellowing and

necrotic spots: Above, new leaves blackened and deformed from SO2

fumigation; Below, closeup.

Control

In Ohio forest nurseries, needle mottling proved to be an excellent characteristic for classifying eastern white pine seedlings as

112
F-522559; F-522839

Figure 32-5.—Above, White ash: chronic foliar symptoms of yellowing and

flecking from SO2 fumigation; Below, sugar maple: acute S0= injury from

fumigation, with intercostal breakdown in tissues.

tolerant or sensitive to ambient pollution. Seedlings that show

needle symptoms of air-pollution injury can then be eliminated

during normal grading practices. Further selection and breeding of tolerant forest tree species should be encouraged.

113
IV. STORAGE MOLDS

DRAKE HOCKING*

Molds of many species may attack foliage or roots of stored

seedlings, causing mortality, deformation, and stunting. Usually,

losses from molding are a result of careless handling of stock or

poor storage facilities.

Molding of coniferous foliage may cause severe losses by reducing survival and impairing first-year growth and form. It is sustained and encouraged by high temperature, high humidity, and

free water. Water on foliage might originate by lifting stock during rain, wetting during packaging, use of excessively wet packaging materials, or condensation due to fluctuating storage temperatures.

Critical temperatures to prevent molding have not been established, but progressively lower temperatures at least down to 0° C

result in progressively less molding. Seedlings stored at subfreezing temperatures generally do not mold, nor do seedlings heeledin in snow. The hazard of molding may increase, however, during

the thawing-out period at the end of storage. Danger and severity

of molding tend to increase with duration of storage, if conditions

conducive to molding persist.

Tree species differ in their susceptibility to molding, perhaps

due to physiological differences or to morphological ones, such as

better ventilation due to longer or stiffer needles.

Fungi that cause molding have seldom been identified, and only

little studied. Genera mentioned as important are Botrytis, Fusarium, Rhizoctonia, Cylindrocarpon, Phacidium, Pythium, Epicoccnniy Phoma, and Pénicillium, Some authors report finding sterile

mycelia.

Inoculum is commonly present on soil adhering to roots and

foliage or in packing materials. Thirty-seven fungi isolated from

moldy seedlings commonly occur in nursery soils ; 10 isolated with

significant frequency developed best under similar temperature

and moisture conditions. Thus, storage conditions may be a more

important factor in molding than the presence of any specific

fungus or group of fungi.

Fungicides have been recommended to control molding during

storage, but experimental testing generally provides negative or

inconclusive results. Packing with peat has been found more

* Research Scientist, Canadian Forestry Service, Northern Forest Research

Centre, Edmonton, Alberta, Can.

114
effective than applying fungicides for mold control, and shingletow packing is more inhibitory than peat.

Hazard of root rotting has been reported serious among several

deciduous species, the mortality reaching 100 percent among stock

heeled-in in shingle tow. Some mortality may have resulted from

toxins in the shingle tow, however.

An unknown agent, probably an infectious microorganism,

caused a storage breakdown of hybrid poplar cuttings; prestorage fungicidal dips gave satisfactory control.

Selected References

Adams, Ronald S., Samuel F. Gossard, and John R. Ritchey.

1967. Phytoactin does not improve survival of stored Monterey pine and

Douglas-fir seedlings. Tree Planters' Notes 18(4) :8--10.

Brown, R. M.

1971. Cold storage of forest plants: A review. Q. J. For. 65, p. 305-315.

Carlson, L. W, and C. Lindquist.

1968 Poplar hybrid storage breakdown; unknown cause. Fungicide and

nematicide tests. Results of 1968; p. 93. Am. Phytopath. Soc. Washington, D. C.

Cram, W. H., A. C. Thompson, and T. Elliott.

1967. Nursery cultural investigations. Storage studies. Tree Nursery,

PFRA, Indian Head, Sask., Can., Summ. Rep. 1966:23-25.

Deifenbacher, Forrest W., and Ernest Wright.

1954. Refrigerated storage of conifer seedlings in the Pacific Northwest.

J. For. 52:936-938.

Hellmers, Henry.

1962. Physiological changes in stored pine seedlings. Tree Planters' Notes

No. 53, p. 9-10.

Hocking, D.

1971. Effect and characteristics of pathogens on foliage and buds of coldstored white spruce and lodgepole pine seedlings. Can. J. For. Res.

1:208-215.

Hocking, Drake, and Ralph D. Nyland.

1971. Cold storage of coniferous seedlings: A review. AFRI Res. Rep.

6, 70 p. State Univ., Coll. For., Syracuse, N.Y.

Krueger, Kenneth W.

1968. Investigations of shingle tow packing material for conifer seedlings. USDA For. Serv. Res. Pap. PNW-63, 10 p. Pacific Northwest

For. and Range Exp. Stn., Portland, Oreg.

Lindquist, C. H.

1971. Plant storage studies. Tree Nursery, PFRA, Indian Head, Sask.,

Can., Summ. Rep. 1970:20-21.

Lindquist, C. H.

1972. Plant storage studies. Tree Nursery, PFRA, Indian Head, Sask.,

Can., Summ. Rep. 1971:27-29.

Nyland, R. D.

1970. A progress report about overwinter cold storage of conifers in

New York State. Northeast. Area State Priv. For., Northeast. Area

Nurserymen's Conf., Proc. 1970:39-44.

Thomann, J.

1971. Planting at high altitudes. Foret 24(4) :106.

Vaartaja, 0.

1955. Storage molding of maple seedlings. Can. Dep. Agrie. For. Biol.

Div., Bimon. Prog. Rep. 11(1) :3.

Young, George Y.

1942. Root rots in storage of deciduous nursery stock and their control.

Phytopathology 33:656-665.

115
GLOSSARY

ABIOTIC—of or pertaining to the nonliving. Syn. inanimate.

ACERVULUS—a small subcuticular or subepidermal cushionlike

asexual fruiting body, without a covering of fungus tissue, producing conidia in a moist mass which escapes through a break

in the host tissue.

ACUTE—having a sudden onset and a sharp rise within a short

period of time.

AECIOSPORE—one of several kinds of spores prodirced by a

rust fungus. Formed in and released from a fruiting structure

called an aecium.

ALTERNATE HOST—one or the other of the two unlike host

plants parasitized by a heteroecious fungus such as a typical

rust fungus, i.e., either the white pine or gooseberry host of the

white pine blister rust fungus.

ANTHRACNOSE—a type of plant disease which typically is a

leaf and twig blight. Common on many hardwoods.

APOTHECIUM(IA)—a cup or saucerlike sexual fruiting body

which produces ascospores.

ASCOGENOUS STAGE—the ascospore producing stage of an

Ascomycete.

ASCOMYCETE—a large group of fungi which are characterized

by the free cell formation of spores, usually eight in number, in

a saclike structure called an ascus.

ASCOSPORE—a spore produced in the sexual or perfect fruiting

body of an Ascomycete.

ASEXUAL STAGE—either a vegetative stage or a reproductive

stage in the life cycle of a fungus in which nuclear fusion is

absent and in which reproductive spores are produced by mitosis

or simple nuclear division. Syn. imperfect stage.

AUTOECIOUS—pertaining to a fungus which completes its life

cycle on one host.

BASIDIOMYCETE—a large group of fungi which are characterized by the production of spores, usually four, on a basidium.

BASIDIOSPORE—the spore produced by the sexual stage of the

Basidiomycetes.

BASIDIUM—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.

BIOCIDE—a wide spectrum poison which kills a great number

and variety of organisms.

BIOTIC—of or pertaining to living organisms. Syn. animate.

BLIGHT—a general term for a plant disease causing rapid death

or dieback.

BROOM—an abnormally dense mass of host branches and foliage

in which the typical host growth pattern is lost.

116
CAMBIUM—^the layer of cells that lies between and gives rise bycell division to the secondary xylem (wood) and the secondary

phloem (inner bark).

CANKER—a definite relatively localized necrotic lesion primarily

of the bark and cambium.

CHLAMYDOSPORE—a thick-walled asexual resting spore typically formed by many soilborne fungi.

CHLOROSIS—an abnormal yellowing of the foilage.

CHLOROTIC—abnormally yellow.

CHRONIC—pertaining to a condition which is of long duration.

COLONIZE—^to establish an infection within a host or part of a

host.

CONIDIA—an asexual spore of a fungus, typically produced terminally on a specialized hyphae termed a conidiophore.

CONIDIOPHORE—a specialized hyphae which produces asexual

spores called conidia.

CORTEX—^the primary tissue of a first-year stem or root found

between the epidermis and the primary vascular bundle.

COVER CROP—a crop, natural or introduced, that is grown

alternately with the main crop. Used to prevent erosion and improve soil characteristics.

CULL—a seedling which is rejected because it does not meet

certain specifications.

CULTURAL PRACTICES—a general term for those routine nursery operations required to help seedling growth, i.e., plowing,

watering, weeding, etc.

CUNÉATE—thinner at one end than the other ; wedge shaped.

DAMPING-OFF—^the killing of the seedling by microorganisms

before emergence or the collapse of the seedling stem at ground

level after emergence.

DECAY—the decomposition of plant tissue by fungi and other

microorganisms.

DECLINE—the gradual reduction in health and vigor as a tree is

in the process of slowly dying.

DIEBACK the progressive dying of stems and branches from the

tip downward.

DISEASE—unfavorable change of the function or form of a

plant from normal, caused by a pathogenic agent or unfavorable environment.

ECHINULATE—having many small spines or prickles.

ECTOPARASITE—a parasite (in particular, a nematode) which

lives outside its host.

ENDEMIC—^native to the country or region.

ENDOPARASITE—a parasite (in particular, a nematode) which

lives inside its host.

EPIDEMIC—pertaining to a disease which has built up rapidly

and reached injurious levels.

EPIDERMIS—the outermost layer of cells on the primary plant

body.

117
EXOTIC—introduced from another country or area.

EXÚDATE—matter which oozes out or is secreted out.

FACULTATIVE SAPROPHYTE—an organism which is normally

parasitic but which is capable of living as a saprophyte.

FALLOW—cultivated land allowed to lie idle or unplanted during

the growing season.

FLACCID—deficient in turgor, limp, or flabby.

FLAG—a conspicuous dead branch with the foliage remaining on.

FRUITING BODY—any of a number of kinds of reproductive

structures which produce spores.

FUMIGATION—to apply vapor or gas to, especially for the purpose of disinfecting or destroying pests.

FUNGICIDE—chemical which is toxic to fungi.

FUNGI IMPERFECTI—SL grouping of miscellaneous fungi which

lack a known sexual stage and which are classified, therefore,

according to the characteristics of their asexual stages.

GALL—a pronounced swelling on a woody plant caused by certain

fungi, bacteria, insects, or nematodes.

GENICULATE—bent abruptly at an angle, like a knee.

GERMINATE—to begin growth from a seed or spore.

GIRDLE—to destroy or remove the tissue, particularly living

tissue in a rough ring around a stem branch or root.

HETEROECIOUS—pertaining to a fungus which must pass a

part of its life cycle on each of two different unrelated hosts, i.e.,

some rust fungi.

HOST—^the plant on or in which a pathogen exists.

HOST RANGE—all hosts which a particular pathogen attacks.

HOST-SPECIFIC—a term used to describe those pathogens which

attack only certain species of hosts.

HYALINE—transparent, having no color.

HYPHA—one of the filamentous threads which make up the

fungus body.

HYPOCOTYL—that part of the axis of a developing embryo just

below the cotyledons.

HYSTEROTHECIUM—a specialized fruiting body of needle cast

fungi which produces ascospores, is usually elongate, covered,

and opens at maturity by a long slit.

IMPERFECT STAGE—that part of the life cycle of a fungi in

which only conidia and no sexual spores are produced. Syn.

asexual.

INCITE—to cause a disease.

INDIGENOUS—native to a particular region or environment.

INFECT—to invade and cause a disease.

INFEST—to be present within an area in such numbers as to

be a disease hazard.

INOCULATE—to place a pathogen on or in a host in a position

in which it is capable of causing a disease.

INOCULUM—the spores, mycelium, sclerotia, or other propagules

of a pathogen which initially infect a host or crop.

118
INTERCELLULAR—lying or growing between the cells.

INTRACELLULAR—lying or growing within the cells.

LATENT INFECTION—an established infection which does not

show its presence.

LEAF SPOT—a leaf disease characterized by numerous distinct

lesions.

LESION—a defined necrotic area.

MACROCONIDIA—the larger of two types of conidia produced

by certain fungi, such as Fnsarium spp.

MICROCONIDIA—the smaller of the two types of conidia produced by certain fungi.

MICROSCLEROTIUM—a very small (microscopic) sclerotium.

MORIBUND—being in a dying state.

MYCELIUM—a mass of hyphae which forms the vegetative filamentous body of a fungus.

NECROSIS—death of plant cells usually resulting in darkening

of the tissue.

OOSPORE—the sexually produced resting spore of the water

molds.

PATHOGEN—an organism which causes a disease.

PATHOGENIC—capable of causing a disease.

PARASITE—an organism living on and nourished by another

living organism.

PERFECT STAGE—the stage in which the sexual spore stage is

produced. Syn. sexual stage.

PERITHECIUM—a closed flasklike sexual fruiting body formed

by certain Ascomycetes in which ascospores are produced.

PHELLUM—the suberized tissue produced by the cork cambium

in the bark.

PHELLODERM—the parenchjmiatous type of tissue produced to

the inside by the cork cambium.

PHLOEM—the tissues of the inner bark responsible for the transport of elaborate foodstuffs.

PHOTOLYTIC—pertaining to the chemical decomposition due to

the action of radiant energy (sunlight).

PHYCOMYCETE—a group of lower fungi which includes the

water molds.

PHYTOTOXIC—a chemical which is toxic to plants.

PYCNIDIUM—an asexual type of fruiting body, typically flask

shaped, in which asexual spores or conidia are produced.

PYCNIDIOSPORE—an asexual spore or conidium produced

within a pycnidium.

PYCNIOSPORE—a specialized spore produced in a pycnium by

the rust fungi.

RESISTANT—able to withstand without serious injury, attack

by an organism, or damage by a nonliving agency but not

immune from such attacks.

ROOT CROWN—the uppermost portion of the root system where

the major roots join together at the base of the stem.

ROT—see decay.

119
SAPROPHYTE—an organism using dead organic material as

food.

SCLEROTIUM—a firm, frequently rounded multicellular resting

structure produced by fungi.

SEPTATE—having cross walls which divide hyphae or spores

into a number of separate cells.

SEPTUM—the cross wall which divides a hypha or spore into two

or more distinct cells.

SEXUAL STAGE—the stage in the life cycle of a fungus in which

spores are produced after sexual fusion. Syn. perfect stage.

SPORE—the reproductive structure of the fungi and other lower

plants.

SPORULATE—to produce and release spores.

SPREADER—a chemical additive used in the preparation of

fungicide sprays to improve the distribution of the spray on the

plant foliage.

STICKER—a chemical additive which is added to fungicide sprays

to improve the fungicide's retention on the plant surface.

STOMA, pi. STOMATA—a pore in the leaf epidermis, surrounded

by two guard cells, leading into an intercellular space within

the plant.

STROMA, pi. STROMATA—a cushionlike body on or in which

fungus fruiting bodies are formed.

SUSCEPTIBLE—unable to withstand attack by an organism or

damage by a nonliving agency without serious injury.

SYMPTOM—the evidence of disturbance in the normal development and function of a host plant, i.e., chlorosis, necrosis, galls,

brooms, stunting, etc.

SYSTEMIC—affecting or distributed throughout the whole plant

body.

TAPROOT—the primary descending root of a plant off of which

the secondary or lateral roots branch.

TELIOSPORE—the spore of the rust fungi from which the perfect stage of the basidium and basidiospore arise.

TELIUM—an aggregation of teliospores of the rust fungi.

TRANSLOCATE—the transfer of elaborated food materials within the plant.

UREDIOSPORES—one of the many spore stages produced by the

rust fungi in their complicated life cycle. These spores are

produced in a fruiting body called a uredium.

UREDIUM—one of the many types of fruiting bodies formed by

the rusts in their complicated life cycle. Urediospores are formed

in this fruiting body.

XYLEM—the woody conducting tissues of the stem and root.

WILT—Si type of plant disease characterized by the sudden loss

in turgor and collapse of the succulent parts of the affected

plants.

ZOOSPORE—a motile free swimming spore produced by the

water molds.

120
INDEX TO HOST TREES, WITH SCIENTIFIC EQUIVALENTS

Host Tree Article Number

Arborvitae, Oriental, Thuja orientalis L. 23

Ash, Fraxinus spp. 1

green, F. pennsylvanica Marsh. 1

Aspen, Populus spp. 28, 29, 30

bigtooth, P. grandidentata Michx. 28

quaking, P. tremuloides Michx. 28, 29

Birch, Betulsi spp. 8

Butternut, Juglans cinérea L. 27

Catalpa, northern. Catalpa speciosa Warder 1

Chestnut, Castanea spp. 6, 15

Chinkapin, Castanea spp. 15

Chinkapin, Castanopsis spp. 15

Cottonwood, Populus spp. 8, 28, 29, 30

eastern, P. deltoides Bartr. 28, 29, 30

black, P. trichocarpa Torr. & Gray 28

Jacks, Populus X jackii Sarg. 30

narrowleaf, P. angustifolia James 28

plains, P. sargenta Dode 29

Cypress, CupressUrS spp. 8, 13, 22, 23

Arizona, C. arizonica Greene 23

Italian, C sempervirens L. 13, 23

Monterey, C macrocarpa Hartw. 13, 23

Dogwood, Cornue spp. 9

Douglas-fir, Pseudotsuga menziesii (Mirb.)

Franco 3, 7, 11, 14, 24, 25, 29

Elm, Ulnvus spp. 1

American, U, americana L. 1

Siberian, U, pumila L. 1

Fir, Ahies spp. 3, 7, 8, 25

balsam, A. balsamea (L.) Mill. 7, 25

Fraser, A. fraseri (Pursh) Poir. 5, 7

grand, A. grandis (Dougl.) Lindl. 25

red, A. magnifica A. Murr. 3

subalpine, A. lasiocarpa (Hook.) Nutt. 25

white, A. concolor (Gord. & Glend.) Lindl. 3, 25

Hackberry, Celtis spp. 1

Hemlock, Tsuga spp. 11, 12, 24

eastern, T. canadensis (L.) Carr. 24

western, T. keterophylla (Raf.) Sarg. 11, 12

Honeylocust, Gleditsia triacanthos L. 1

Incense-cedar, Lihocedrus spp. 8

121
Juniper, Juniperus spp. 8, 22

Rocky Mountain, J. scopulorum Sarg. 22

Larch, Larix spp. 29

Locust, black. Robinia pseudoacacia L. 1

Maple, Norway, Acer platanoides L. 8

Mulberry, Russian, Morus alba f. tartárica Seringe. 1

Oak, Querctcs spp. 1, 6, 15, 17

Pine, Pinus spp 2, 3, 4, 5, 7, 9, 12, 14, 15, 16,

17, 18, 19, 20, 21, 25, 29

Austrian, P. nigra Arnold 7, 19, 21

bishop, P, muricata D. Don 16

Coulter, P. coulteri D. Don 12, 16

Digger, P. sabiniana Dougl. 16

eastern white, P. strobu^ L. 7, 14, 18, 19

foxtail, P. balfouriana Grev. & Balf. 18

jack, P. banksiana Lamb. 7, 14, 16, 17, 19, 20

Jeffrey, P. jeffreyi Grev. & Balf. 3, 12, 16, 17, 19

knobcone, P. attenuata Lemm. 16

limber, P. flexilis James 18, 25

loblolly, P. taedaL. 4, 15, 17, 19, 20, 31

lodgepole, P. contorta Dougl. 12, 14, 16, 17, 19, 29

longleaf, P. palustris Mill. 4, 15, 19, 20

Monterey, P. radiaba D. Don 3, 16, 19, 21, 29

Mugo, P. mugo Turra 7, 19

pitch, P. rigida Mill. 15, 17, 19

pond, P. serótina Michx. 4, 15

ponderosa, P. ponderosa Laws. 3, 12, 16, 17, 19, 25, 29

red, P. resinosa Ait. 5, 7, 12, 14, 19

sand, P. c^wsa (Chapm.) Vasey 4, 9

Scotch, P. sylvestris L. 7, 14, 16, 17, 19, 20, 29

shortleaf, P. echinata Mill. 4, 15, 17, 19

slash, P. elliottii var. elliottii Engelm. 4, 15, 19, 20, 31

spruce, P. glabra Walt 4

sugar, P. lambertiana Dougl. 3, 12, 18

Virginia, P. virginiana Mill. 4, 17

western white, P. monticola Dougl. 18, 19

whitebark, P. albicaulis Engelm. 18

Flanetreey London, Platanus acerifolia (Ait.) Willd. _ 26

Poplar, Populus spp. 28, 29, 30

balsam, P. balsamifera L. 28, 29

black, P. nigra L. 28

Carolina, P. canadensis Moench 28

Rio Grande, P. fremontii var. wislizenii S. Wats. 29

white, P. dZ&a L. 28

Prunus, Prunus spp. 8

Redcedar, eastern, Juniperus virginiana L. 8, 9, 22, 23

Redwood, Sequoia sempervirens (D. Don) Endl. 13

Russian-olive, Elaeagnu^ angustifolia L. 1

Sequoia, giant, Sequoiadendron giganteum (Lindl.)

Buchholz 3, 13, 23

122
Spruce, Picea spp. 7, 12, 14, 25

hlsick, P. mariana (Mill.) B.S.P. 7, 14, 25

blue, P. pungens Engelm. 7, 25

Engelmann, P engelmannii Parry 25

Norway, P. abies Karst. 7, 12, 14, 25

red, P. rubens Sarg. 12, 25

Sitka, P. sííc/^e725Ís (Bong.) Carr. 24

white, P. glauca (Moench) Voss 7, 14, 25

Sweetgum, Liquidambar styraciflvu L. 1

Sycamore, Platanus spp. 1, 26

American, P. occidentalis L. 26

California, P. racemosa Nutt. 26

Tanoak, Lithocarpits densiflorus (Hook. & Arn.) Rehd. 15

Thuja, Thuja spp. 22

Walnut, Juglans spp. 5, 6, 7, 27

black, /. nigra L. 5, 6, 7, 27

California, /. californica S. Wats. 27

Hinds, /. hindsii Jeps. 27

Persian, /. regia L. 27

White cedar, Chamaecyparis spp. 22

Willow, Salix spp. 8

Yellow-poplar, Liriodendron tulipifera L. 1, 5, 7

INDEX TO DISEASES

Article No.

Disease

A

Agrobacterium tumefaciens 8

Air pollutant damage 32

Ascochyta piniperda 12

B

Black root rot 4

Botrytis cinérea 13

Brown spot needle blight 20

c

Cercospora blight 23

Cercospora sequoiae 23

Charcoal Root Disease 3

Chlorosis 31

Comandra rust 17

Cronartium coleosporioides 17

Cronartium comandrae 17

Cronartium comptoniae 17

Cronartium funiforme 15

Cronartium quercuum 17

123
Cronartium ribicola 18

Crown gall8

Cylindrocladium floridanum7

Cylindrocladium root rot7

Cylindrocladium scoparium7

Cytospora spp.30

D

Damping-off1

Diaporthe lokoyae 11

Dothistroma needle blight21

Dothistroma pini21

E

Eastern gall rust 17

Endocronartium harknessii 16 F

Fusarium oxysporum2

Fusarium Root Disease2

Fusarium spp.30

G

Gloeosporium platani26

Gnomonia leptostyla27

Gnomonia platani26

Grey Mold 13 H

Hemiphacidium planum 25 L

Lecanosticta acicola20

Lophophacidium hyperboreum25

Lophodermium needle cast 19

Lophodermium pinastri 19 M

MacrophowÂna pha^eoli 3,4

Marssonina juglandis 27

Marssonina leaf blight 28

Marssonina populi28

Melampsora medusae29

Melampsora rust 29

Mycorrhizae 10 N

Nematodes9

P

Peridermium harknessii 16

Phacidium abietis 25

Phomopsis blight 22

Phomopsis canker 11

Phomopsis juniperovora 22

Phomopsis lokoyae 11

Phomopsis spp. 30

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Phytophthora cactorum 6

Phytophthora cinnamomi 5, 6

Phytophthora citricola 6

Phytophthora root rot 5, 6

R

Rosellinia herpotrichoides 24

Rosellinia needle blight 24

s

Sarcotrichila balsameae 25

SarcotricMla piniperda 25

Scirrhia acicola 20

Scirrhia pint 21

Sclerotinia fuckeliana 13

Sclerotium bataticola 3, 4

Scleroderris canker 14

Scleroderris lagerbergii 14

Septoria leaf spot and canker 30

Septoria rmisiva 30

Sirococcus strobilinus 12

Sirococcus tip blight 12

Snow blight 25

Southern fusiform rust 15

Stalactiform rust 17

Storage molds IV

Sweetfern rust 17

Sycamore anthracnose 26

Systremma acicola 20

w

Walnut anthracnose 27

Western gall rust 16, 17

White pine blister rust 18

Y

Yellows 31

^ U.S. GOVERNMENT PRINTING OFFICE: 1975 0—551-594

125