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Contents

5.2.1 Introduction 103

5.2.1.1 What are mycorrhizae? 103

5.2.1.2 Types of mycorrhizae 104

Ectomycorrhizae 104

Ectendomycorrhizae 110

Vesicular-arbuscular mycorrhizae 110

5.2.1.3 Major benefits of mycorrhizae 115

5.2.2 Current Status of Mycorrhizae in

Container Nurseries 117

5.2.2.1 Results of nursery survey 117

5.2.2.2 Observations in Pacific Northwest 

nurseries 121

5.2.2.3 Mycorrhizae: why some seedlings are 

mycorrhizal and some are not 122

5.2.3 How to Check Seedlings for

Mycorrhizae 123

5.2.3.1 Ectomycorrhizae 123

5.2.3.2 Ectendomycorrhizae 124

5.2.3.3 Vesicular-arbuscular mycorrhizae124

5.2.4 Mycorrhizal Fungi That Fruit in

Container Nurseries 125

5.2.5 Determining the Need for Mycorrhizal

inoculation 127

5.2.5.1 In-nursery benefits 127

5.2.5.2 Outplanting benefits 127

5.2.6 Sources of Inoculum and Inoculation

Techniques 130

5.2.6.1 Soil inoculum 130

5.2.6.2 Spore inoculum 130

5.2.6.3 Mycelial inoculum 135

5.2.6.4 Vesicular-arbuscular inoculum 137

5.2.6.5 Isolate selection and ecotypic variation 139

5.2.7 Evaluating Inoculation Success 141

5.2.7.1 Rating mycorrhizal formation 149

5.2.7.2 Designing outplanting trials 149

5.2.7.3 Economic considerations 150

5.2.8 Factors Affecting Mycorrhizal

Development 152

5.2.8.1 Root development 152

5.2.8.2 Fertilizer 152

5.2.8.3 Water 154

5.2.8.4 Growing media 155

5.2.8.5 Temperature 156

5.2.8.6 Pesticides 157

Sterilants 157

Fungicides 157

Herbicides 158

Insecticides and nematicides 158

General pesticide recommendations 158

5.2.9 Conclusions and

Recommendations 159

5.2.10 References 160

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Although considerable research is currently in progress on the role 

of mycorrhizae in plant nutrition and practi cal uses in forestry, an 

abundance of information and concepts are available for 

immediate use in tree seedling nurseries. Our objectives for this 

chapter are fourfold:

1. Describe the different types of mycorrhizae common to 

forest tree seedlings grown in container nurseries.

2. Define the benefits imparted by mycorrhizae to

seedling nutrition, growth, and survival.

3. Document the occurrence of mycorrhizae on container grown 

seedlings and describe how routine nursery practices affect 

the development of mycorrhizae.

4. Recommend ways for nursery managers to incorporate 

mycorrhizal management into their cultural regimes and offer 

management strategies to enhance mycorrhizal development 

and subsequent seedling survival and growth after 

outplanting.

5.2.1.1 What are mycorrhizae?

The word mycorrhizae literally means "fungus roots" and defines 

the intimate associations between plant roots and specialized soil 

fungi, the mycorrhizal fungi. Nearly all the world's land plants form 

some type of mycorrhiza, and with few exceptions, all major forest 

tree species form mycorrhizae. Two major mycorrhizal types 

prevail among forest trees: ectomycorrhizae, which are formed 

with the important coniferous species of the Pinaceae and 

hardwoods in the Fagaceae and Betulaceae; and 

vesicular-arbuscular (VA) mycorrhizae, which are common on 

other hardwoods, particularly in the maples, sweetgums, cedars, 

and redwoods. Although similar in overall function and benefit to 

the host plant, these two types of mycorrhizae differ strongly in 

regard to the fungi involved, their morphology, and potential 

applications in forest tree nurseries. Table 5.2.1 lists the major 

genera of forest trees raised in nurseries of temperate North 

America along with the types of mycorrhizae they form.

First we will describe each major type and how to identify 

them and then outline the major benefits.

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5.2.1 Introduction

A major advantage in rearing container seedlings is the production 

of large, robust seedlings within a single growing season. This 

contrasts with the 2- to 3-year cycle of producing plantable 

seedlings of desired size in bareroot seedling nurseries in the 

western and northern United States. Most current criteria of 

seedling quality are limited to the condition and size of the seedling 

stem and foliage. Less attention is paid to the quality of roots on 

nursery seedlings, even though we are well aware of the 

paramount importance of roots in providing structural support and 

nutrient and water uptake. Thus, to completely evaluate the 

"health" of a seedling and predict its survival potential, we must 

increase our awareness of root quality.

To develop criteria for evaluating seedling root quality in the 

nursery, we must incorporate knowledge of the root dynamics of 

wild seedlings. This knowledge is critically important because once 

seedlings are removed from the nursery and planted into soil, the 

roots must function under soil conditions as mediated by complex 

and uncontrolled environmental and biotic factors. These soil 

conditions will differ drastically from the well-watered, well-fertilized 

nursery growing media.

In natural soils, all forest trees form symbiotic, mutually beneficial 

associations between their roots and specialized soil fungi. The 

fungus-root organ is called a mycorrhiza (mycorrhizae is plural). 

Mycorrhizae provide many benefits to the seedling and adult tree, 

especially in enhancing water and nutrient uptake. Indeed, 

seedlings strongly depend upon mycorrhizae for growth and 

survival as evidenced by the failure of nonmycorrhizal seedlings to 

survive when planted into soil lacking mycorrhizal fungi (Trappe 

1977). Thus, the presence and abundance of mycorrhizae must be 

a major consideration for evaluating root system health and 

predicting outplanting performance.
5.2.1.2 Types of mycorrhizae

Ectomycorrhizae. Ectomycorrhizae develop on the short, feeder 

roots, as opposed to the longer, woody, structural lateral roots. In 

fact, once a root develops a lateral meristem and starts forming 

woody tissue, mycorrhizae can no longer form. Ectomycorrhizae 

can be easily recognized by the characteristic fungal sheath or 

mantle tissue that envelopes the feeder roots; often the fungal 

mycelium, or thread-like mold growth, can be seen emanating 

directly from the mantle and colonizing the soil or rooting substrate 

(fig. 5.2.1 and 5.2.2). When an ectomycorrhiza is sectioned and its 

internal anatomy is examined under a microscope, we can see the 

second major characteristic of ectomycorrhizae: the intercellular 

growth of the fungus between the epidermal and cortical cells that 

forms the Hartig net (fig. 5.2.3). It is within this extensive zone of 

fungus-root cell contact that nutrients and water are exchanged 

between fungus and host; the fungus brings in and releases to the 

host nutrients and water and in return receives plant-made sugars 

and other products of photosynthesis.

The fungi that form ectomycorrhizae are primarily Basidiomycotina 

and Ascomycotina (table 5.2.2), including many of the common 

forest mushrooms (fig. 5.2.4 and 5.2.5) and puffballs (fig. 5.2.6), as 

well as the hypogeous (below-ground) fruiting fungi called truffles 

(fig. 5.2.7-5.2.9). Well-known fungal genera that form 

ectomycorrhizae include Amanita, Boletus, Hebeloma, Laccaria, 

Lactarius, Pisolithus, Rhizopogon, Russula, Scleroderma, Suillus, 

and Tricholoma (all Basidiomycotina), and Cenococcum and Tuber

(Ascomycotina) (see Miller (1982) for a complete listing of 

ectomycorrhizal fungus genera). Another common ectomycorrhizal 

fungus in seedling nurseries is Thelephora terrestris (and closely 

related species in the same genus). Thelephora fruiting bodies 

(or sporocarps) commonly occur as leathery, erect brown sheets 

or mats on the bases of seedling stems (fig. 5.2.10) or on and 

around the drainage holes of individual containers or bottoms of 

Styroblocks® (fig. 5.2.11 and 5.2.12). Thelephora species are the 

most common ectomycorrhizal fungi in container nurseries; we will 

discuss their occurrence and importance in later sections.

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Because most container tree seedling nurseries use artificial 

growing media, which lack ectomycorrhizal fungi, it is important to 

understand how container seedlings can become ectomycorrhizal, 

either naturally or by controlled methods. Many ectomycorrhizal 

fungi produce spores that are disseminated by the wind, allowing 

long-distance dispersal from forests to nurseries for inoculation of 

seedlings. However, the farther a nursery is located from 

ectomycorrhizal forests or large tracts of ectomycorrhizal trees, the 

less will be its chance of receiving natural wind-dispersed spore 

inoculum. Within the nursery, fruiting bodies produced on 

ectomycorrhizal seedlings offer a reliable source of spores to 

inoculate neighboring seedlings and future crops. The practical 

implications of spores as sources of inoculation will be discussed in 

detail in section 5.2.6.2.

The structural appearance of ectomycorrhizae is a function of both 

fungus and host plant. Thousands of different fungi form 

ectomycorrhizae, many with more than one host plant, so the 

overall appearance of different fungus-host combinations can vary 

tremendously. Figures 5.2.13 through 5.2.16 illustrate 

ectomycorrhizal form and diversity. Ectomycorrhizal morphology is 

often characteristic for a particular host genus. For example, the 

root tips of ectomycorrhizae in pines often branch dichotomously 

into complex structures (fig. 5.2.17). Other ectomycorrhizal forms 

range from simple cylinders to complex, pinnate, coralloid, or even 

compact tubercle forms (fig. 5.2.18). The amount of mycelium 

emanating from an ectomycorrhiza is another important diagnostic 

character. External mycelium (or hyphae) can range from sparse, 

nearly invisible threads to prolific wefts and root-like strands of 

hyphae (rhizornorphs) that transport nutrients and water (fig. 

5.2.15 and 5.2.18).

Ectendomycorrhizae. Ectendomycorrhizae represent a second 

type of mycorrhiza, which can be abundant on nursery stock, 

particularly pines and spruces. Ectendomycorrhizae look like 

ectomycorrhizae in general form but usually lack the thick, often 

colorful mantle and abundant visible external hyphae usual for 

ectomycorrhizae (fig. 5.2.19). In cross section, the fungus can be 

seen penetrating into cortical cells as well as forming a Hartig net 

between them (fig. 5.2.20). Although we know little of the ecology 

of ectendomycorrhizal fungi or their effects on seedling nutrition, 

growth, and survival, ectendomycorrhizae have been shown to be 

beneficial in some instances (LoBuglio and Wilcox 1987, Wilcox 

and GanmoreNeumann 1974). The fungi are Ascomycotina and 

mostly lack mushroom-like fruiting structures, although some form 

small cup-shaped fruiting bodies on the surface of the growing 

medium.

Vesicular-arbuscular mycorrhizae. Vesicular-arbuscular (VA) 

mycorrhizae appear strikingly different from ectomycorrhizae: they 

do not modify root morphology and the fungal component is 

invisible to the unaided eye. Roots must be differentially stained 

and observed under the microscope to satisfactorily discern the 

fungal structures and degree of root colonization (fig. 5.2.21). As 

implied in the name, two structures characterize the VA 

mycorrhiza-vesicles and arbuscules. Vesicles are balloon-shaped 

structures, usually filled with lipids (oil droplets), that serve both as 

energy storage organs and as reproductive structures (fig. 5.2.22). 

Arbuscules are finely branched, intracellular, short-lived structures 

that serve as nutrient exchange sites between fungus and host (fig. 

5.2.23). VA mycorrhizae also have abundant fungal mycelium that 

ramifies through the root cortex and extends out into the soil.

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Zygomycotous fungi in the family Endogonaceae form VA 

mycorrhizae and number a few hundred species among the 

genera Acaulospora, Entrophospora, Gigaspora, Glomus, 

Sclerocystis, and Scutellospora (table 5.2.2). Unlike the 

mushrooms and puffballs characteristic of ectomycorrhizal fungi, 

VA mycorrhizal fungi usually form relatively large (30 to 900 gm in 

diameter), solitary spores or clumps of spores in the soil (fig. 

5.2.24 and 5.2.25). Because of their size and location, these 

spores are not wind disseminated like the much ,smaller spores of 

ectomycorrhizal fungi. Thus their movement is primarily by 

processes of soil movement; small animals and insects may also 

eat them and disseminate the spores in fecal droppings. This 

restrictive spore dispersal mechanism is significant because it 

greatly reduces their ability to colonize container seedlings growing 

in artificial media, which lack VA mycorrhizal fungi. In section 

5.2.6.4 we will discuss how VA mycorrhiza-forming plants can be 

inoculated.

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5.2.1.3 Major benefits of mycorrhizae

Mycorrhizae benefit plant nutrition, growth, and survival in many 

ways; the best known benefits are enhanced uptake of water and 

mineral nutrients, especially phosphorus and nitrogen (Bowen 

1973). These benefits are due in part to the exploration of soil for 

nutrients and water by hyphae to an extent far beyond the 

capabilities of roots alone. Some researchers estimate that 

mycorrhizal fungus hyphae can explore volumes of soil hundreds 

to thousands of times greater than can roots. Ectomycorrhizal 

fungi also produce growth regulators that stimulate feeder root 

elongation and branching, thus increasing the total number of 

feeder roots produced. Such root branching also benefits 

absorption of nutrients by increasing root surface area. Some 

ectomycorrhizal fungi produce dense mycelial mats in the soil for 

capturing nutrients, while others also produce rhizomorphs--large 

strands of parallel hyphae--that act as conduits for the flow of 

nutrients to and from the ectomycorrhizae (fig. 5.2.26). 

Ectomycorrhizae also reduce root respiration, which would 

increase root longevity (Marshall and Perry 1987). Although VA 

mycorrhizal fungi do not alter gross root morphology, they too 

explore great volumes of soil with their external mycelia and thus 

return nutrients and water from a soil zone beyond the limits of 

root hairs. Readers are referred to the texts by Harley and Smith 

(1983), Marks and Kozlowski (1973), and Schenck (1982) for 

more detailed information concerning mycorrhizal effects on plant 

mineral nutrition.
Mycorrhizal fungi can protect roots against pathogens in several 

ways (Marx 1972). The fungus mantle of ectomycorrhizae provides 

a direct barrier against pathogen entry. Moreover, many 

ectomycorrhizal fungi produce antibiotics antagonistic to some root 

pathogens (Wilkins and Harris 1944, Wilkins and Partridge 1950). 

For example, Marx (1969a, 1969b, 1970) reported strong antibiotic 

production by the ectomycorrhizal fungus Leucopaxillus cerealis 

against Phytophthora cinnamomi. In nursery studies, Laccaria 

laccata suppressed Fusarium oxysporum on Douglas-fir seedlings 

(Sylvia 1983; Sylvia and Sinclair 1983a, 1983b). Unfortunately, 

much of the exploratory research of ectomycorrhizal control of 

pathogens has been done with pure cultures of fungi or in small, 

isolated studies. Use of mycorrhizae for biological control of root 

pathogens is lagging behind other applications and needs serious 

research attention.

Nursery managers should be aware of one other aspect of 

mycorrhiza--disease interactions: mycorrhizae indirectly prote ct 

plants against many types of pathogens (Schenck 1981) by 

benefiting plant growth. Healthy plants with well-balanced nutrition 

resist disease better than plants with poor nutrition. Mycorrhizae 

contribute vitally to adequate plant nutrition: they thereby contribute 

indirectly to the plant's resistance to disease. Because timing may 

be critical for resistance, the sooner the mycorrhizal fungus is 

present in the substrate, the greater the potential for pathogen 

control. By ensuring that mycorrhizae develop on seedlings, ery 

managers also provide some degree of protect against pathogens.

Other benefits of mycorrhizae include enhanced ro ing of 

cuttings (Linderman and Call 1977, Navratil Rochon 1981), 

increased root regeneration, increasE salt tolerance, and 

reduced drought stress (Parke et 1983a). Some of these 

beneficial attributes may be important in nursery management 

for mycorrhizae, whereas others are important for seedling 

survival an growth after outplanting.

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rhizae are important during nursery culture. However, most believe 

that mycorrhizae are most important after the seedlings are 

outplanted. Eighty percent of the managers indicate they can 

recognize mycorrhizae on their seedlings. About two thirds of them 

survey their stock for mycorrhizae but report only low to moderate 

levels of mycorrhizal development. Our observations of some of 

their stock indicate that they likely underestimate the amount of 

mycorrhizae (table 5.2.4). Many managers find fruiting bodies in 

their nurseries but usually cannot identify them. When fruiting 

bodies have been identified by or for the nursery manager,

5.2.2 Current Status of Mycorrhizae in

Container Nurseries

5.2.2.1 Results of nursery survey

To our knowledge, there has never been a systematic survey of the 

types of mycorrhizae found in container tree seedling nurseries. To 

this end, we sent a questionnaire to container tree seedling 

nurseries across the United States and Canada, and 78 nursery 

managers responded (table 5.2.3). Although many believe that it is 

important to inoculate, only 6% of these nurseries have fungal 

inoculation programs. Seventy -seven percent of the nursery 

managers believe that mycorrhizae are important; less than half of 

them think that mycor-
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In addition to the survey questionnaire, we asked the nursery 

managers to send a representative sample of their seedlings to 

our laboratory for evaluation. We examined up to 50 seedlings of 

19 different tree species from 18 nurseries (table 5.2.3). 

Ectomycorrhizal colonization was estimated by type per seedling 

for each nursery. Most seedlings had some ectomycorrhizae; 

many were totally colonized. Thelephora spp. formed the majority 

of ectomycorrhizae among the 19 tree species, especially on 

larch, spruce, and some species of pine. Ectendomycorrhizae of 

undeterminable identity were abundant on several pines, 

especially jack, western white, Austrian, longleaf, ponderosa, and 

Scotch pines. Ectomycorrhizae formed by laurel oak and yellow 

birch were unlike any noted on the conifers (fig. 5.2.29).

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Pisolithus tinctorius (fig. 5.2.27), Laccaria laccata, Thelephora 

terrestris, and Endogone lactiflua (fig. 5.2.28) prove to be the 

most common.
5.2.2.2 Observations in Pacific Northwest nurseries

We have monitored ectomycorrhizal development on container 

seedlings in several Pacific Northwest nurseries for 15 years and 

found that it varies between nurseries and years. However, we 

have noted several consistencies. We see an abundance of conifer 

seedling ectomycorrhizae formed with Thelephora spp. Thelephora 

spp. are well adapted to nursery conditions in which abundant 

water and soluble nutrients stimu late rapid colonization of the 

growing media by the fungus, closely followed by development of 

fruiting bodies (fig. 5.2.10-5.2.12). Another reason for their 

prominence in container nurseries is that Thelephora fruiting bodies 

form early in summer and become a source of spore inoculum for 

the rest of the nursery. Ponderosa pine seedlings often have a high 

degree of colonization by ectendomycorrhizae in addition to 

Thelephora ectomycorrhizae. Engelmann and white spruce are 

typically heavily colonized by Thelephora and Laccaria laccata at 

high rates of soluble and slowrelease fertilizer. However, when 

slow-release fertilizer is eliminated or reduced, Thelephora 

mycorrhizae are readily replaced by E-strain (an unidentified 

Ascomycota that forms ectendomycorrhizae), Amphinema

byssoides (fig. 5.2.30), and occasionally Cenococcum geophilum

mycorrhizae (Hunt 1987). We have also observed the 

ectendomycorrhiza-forming Ascomycotina Sphaerosporella 

brunnea on pines, as have others in Canada (Danielson 1984).

Other trees such as true firs, Douglas-fir, and western hemlock 

often form few or no mycorrhizae despite the same exposure to 

Thelephora and Sphaerosporella spores.

Seedlings in Northwest nurseries frequently have water roots, the 

thick, fleshy, opaque nonmycorrhizal roots lacking roothairs (fig. 

5.2.31) that develop in saturated growing media. We urge nursery 

managers to check seedlings regularly for water roots when 

assessing root quality. Water roots are not colonized by 

mycorrhizal fungi and may even become infected by fungal pathogens (see chapter 1). Water roots are discussed in more detail in 

section 5.2.8.3.

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5.2.2.3 Mycorrhizae: why some seedlings are

mycorrhizal and others are not

Although most container nurseries will have some seedlings 

(especially Douglas-fir and pines) that are ectomycorrhizal with one 

fungus or another, these seedlings (except for seedlings 

ectomycorrhizal with Thelephora) are erratically distributed within 

the nursery. This erratic distribution is caused by the inability of 

ectomycorrhizal fungi to spread vegetatively (that is, with their 

mycelia) from container to container. Each seedling must therefore 

have fungus spores land on and wash into its growing medium, 

find a susceptible feeder root, germinate, and form 

ectomycorrhizae. Thelephora spores do just that with amazing 

efficiency. Thelephora spp. grow rapidly after germination, form 

abundant ectomycorrhizae, and produce fruiting bodies midway 

through the growing season. Such adaptations by Thelephora spp. 

make them the dominant type in container nurseries. Most other 

ectomycorrhizal fungi may not produce fruiting bodies, do not 

distribute their spores through the air, or often grow slowly. Such 

fungi spread even more erratically throughout the nursery than 

Thelephora.
5.2.3 How to Check Seedlings for

Mycorrhizae

Seedlings should be assessed after they have been hardened off. 

During the seedlings' juvenile and rapid growth phases, mineral 

nutrition, especially that of nitrogen, is extremely high. This high 

availability of soluble fertilizer will inhibit most fungi to some extent. 

It is not uncommon to observe proliferation of mycelium and 

mycorrhiza formation as the first stage of seedling hardening 

begins. The timing of mycorrhizal assessment will greatly influence 

the amount of mycorrhizae observed.

Mycorrhizae can be distinguished from pathogenic fungi by 

the presence of visible mycelia surrounding the root and 

the lack of decay.

To assess ectomycorrhizal development, first remove the 

seedling from the container and gently wash the roots free of 

growing media. Suspend the root system in a dish (1 to 2 inches 

deep) that is partially filled with tap water and gently spread the 

root system so that feeder roots are clearly visible. Mycorrhizae 

are then assessed by viewing the immersed roots under a 

stereomicroscope at 5 to 15 times magnification.

5.2.3.1 Ectomycorrhizae

Ectomycorrhizae may be difficult to recognize at first, but with a 

little practice nursery staff can soon distinguish between 

ectomycorrhizal (fig. 5.2.32 and 5.2.33) and nonmycorrhizal (fig. 

5.2.34) feeder roots. Ectomycorrhizae of hardwoods are not as 

easily discernible as those of conifers. The following key characters will guide recognition:

1. Ectomycorrhizae are typically swollen and lack root

hairs.

2. The fungus mantle or sheath is usually a different

color than the feeder roots; some mantles are

brightly colored or pure white (fig. 5.2.32 and

5.2.33).

3. Fungus mycelium or hyphal strands often grow out from the 

mantle tissue, giving a cottony appearance (fig. 5.2.32).

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5.2.3.2 Ectendomycorrhizae

Ectendomycorrhizae are more difficult to recognize. They can 

appear nonmycorrhizal because the fungal mantle can be sparse 

and thin. Ectendomycorrhizae usually lack root hairs, however,

and are usually not significantly swollen. Absolute assessment of 

ectendomycorrhizae or verification of young ectomycorrhizae 

involves examining the roots with a compound light microscope 

for Hartig net formation or intracellular fungus growth. Although 

with training this, too, is easy, nursery staff can usually consult a 

specialist if necessary. Readers are referred to Wilcox (1982) for 

a more detailed description.

Counting total feeder roots during mycorrhizal evaluation is time 

consuming and usually unnecessary. Once you recognize 

ectomycorrhizae, you can easily estimate proportions of the root 

system with ectomycorrhiza colonization by major categories. 

Colonization categories of 1 = none, 2 = low (1 to 25%), 3 = 

medium (26 to 75%), and 4 = high (76 to 100%) will provide functional groupings (Grand and Harvey 1982) to evaluate the 

ectomycorrhizal component of seedling root quality (Benson and 

lyer 1978).

5.2.3.3 Vesicular-arbuscular mycorrhizae

The assessment techniques mentioned in section 5.2.3.2 will not work 

for VA mycorrhizae because roots must be stained and observed under 

a compound light microscope to discern the mycorrhizal structures. If 

nurseries use completely artificial growing media (that is, no soil) to 

grow VA mycorrhizal hosts (maple, sycamore, sweetgum, redwood, 

western redcedar, juniper), few or no plants will have mycorrhizae 

unless inoculated. Remember that VA mycorrhizal fungal spores are 

usually not disseminated through the air. Nursery managers are 

referred to the detailed procedures described by Kormanik and McGraw 

(1982) for staining and assessing VA mycorrhizal roots. Their 

procedures are within a longer text on mycorrhizal methods and 

principles published by the American Phytopathological Society 

(Schenck 1982). We recommend this reference text for nurseries 

developing either ectomycorrhizal or VA mycorrhizal inoculation 

programs.

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4. Mature ectomycorrhizae typically branch several times in 

regular or irregular patterns (fig. 5.2.32 and 5.2.33).

5. Nonmycorrhizal feeder roots are not swollen, are usually 

covered with root hairs, and for many conifer species, are 

unbranched (fig. 5.2.34).

Careful examination of the ectomycorrhizae shown in figures 

5.2.13-5.2.18, 5.2.32 and 5.2.33 will help nursery staff recognize 

these characters.
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5.2.4 Mycorrhizal Fungi That Fruit in

Container Nurseries

Overall ectomycorrhizal diversity is low in container nurseries 

compared to natural conditions. For reasons already mentioned, 

Thelephora spp. are the most common ectomycorrhizal fungi that 

fruit in both container and bareroot nurseries. Fruiting bodies of 

Laccaria laccata (fig. 5.2.35), Inocybe lacera (fig. 5.2.36), Hebeloma crustuhniforme (Castellano and Trappe 1987) and H. arenosa 

(Burdsall and others 1986) (fig. 5.2.37) are next in frequency of 

occurrence, particularly with pines or Douglas-fir. E-strain, 

Amphinema byssoides, and Mycelium radicus atrovirens are 

common on Engelmann spruce grown without slow-release 

fertilizer (Hunt 1987). Occasionally, seedlings (usually hemlocks) 

are colonized by Cenococcum geophilum, which forms a 

characteristic black ectomycorrhizae (fig. 5.2.38). Other fungi have 

been observed but with very low frequency. For example, fruiting 

bodies of Rhizopogon rubescens have been found in ornamental 

stock that was first grown in bareroot beds, then transferred to 

large pots (fig. 5.2.39).
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Nursery managers and foresters should use mycorrhizal 

inoculation as another tool in their effort to grow seedlings and 

reforest land. The effectiveness of inoculation techniques varies by 

host and fungus, so flexibility is paramount to success. One fungus 

(or ecotypic isolate) may accomplish one to many objectives for 

one or many host species (or even seed sources). A flexible 

inoculation program would be able to meet some objectives for one 

portion of the stock and other objectives for other portions of the 

stock. No one fungal species, isolate, or ecotype will meet the 

objectives of all nurseries. The technology is currently available 

to tailor an inoculation program for each nursery, but fine-tuning the 

program to individual nurseries is probably a 2- to 3-year process.

5.2.5.2 Outplanting benefits

The critical test of benefit from mycorrhizal inoculation is 

seedling performance after it is planted in the field (Marx 

1980). Regardless of how mycorrhizal inoculation affects growth in 

the nursery, the seedling must establish and grow once it is 

outplanted. Mycorrhizal inoculation may indeed produce no 

increase in seedling growth in the nursery but will give seedlings a 

better chance to survive or grow better once outplanted 

(Castellano 1987).

5.2.5 Determining the Need for Mycorrhizal

Inoculation

Tree species in the Pinaceae and Fagaceae, which include the 

major coniferous forest species and the oaks, require 

ectomycorrhizae for survival and growth in natural ecosystems. 

This has been convincingly demonstrated in attempts at 

afforestation in the treeless grasslands of the Soviet Union and 

the United States (Mikola 1970). Ectomycorrhizal inoculation has 

proven beneficial in a wide variety of instances, for reclama tion of 

adverse sites, reforestation of clearcut areas, reforestation after 

wildfire, and introduction of exotic species (Marx 1980).

5.2.5.1 In-nursery benefits

Nonmycorrhizal seedlings usually grow well in artificial growing 

media if water and soluble nutrients are supplied. Nonmycorrhizal 

feeder roots of these same seedlings will not properly take up 

water and nutrients from soil after outplanting until they form 

mycorrhizae. The old working premise that "any ectomycorrhizae 

on seedlings are better than none" is under close scrutiny. Some 

ectomycorrhizal fungi are better than others for selected 

applications. We have seen a lag in growth, and a reduction in 

survival, of nonmycorrhizal seedlings and those ectomycorrhizal 

with "nursery-adapted" fungi when outplanted on sites demanding 

quick establishment for survival, for example, droughty, south facing 

steep slopes in the Siskiyou Mountains of southwest Oregon. The 

time needed for seedling root systems to replace nursery-adapted 

fungi with fungi better adapted to site conditions leads to increased 

mortality and reduced initial seedling growth. An effective inoculation program requires mycorrhizal fungi that function efficiently in 

the seedling's growing environment, be it the nursery or in the field. 

The nursery inoculation program must have clear objectives:

1. Reduction in cull percentage in the nursery.

2. Increased stem caliper or leader growth in the nursery and/or field.

3. Protection against pathogens.

4. Rapid mycorrhizal colonization to alleviate

stunting.

5. Increased outplanting survival.
128

Much of the published work on practical application of 

ectomycorrhizal inoculation is concerned with Pisolithus tinctorius

inoculation. Dr. Donald Marx and colleagues at the USDA Forest 

Service Institute for Mycorrhizal Research and Development, 

Athens, Georgia, demonstrated the first wide- scale application of 

ectomycorrhizal inoculation for improving seedling field 

performance. Numerous studies have shown the benefit of P. 

tinctorius ectomycorrhizae to seedling outplanting performance 

(Beckjord and McIntosh 1984; Berry 1982; Dixon et al. 1981; Dixon 

et al. 1984b; Kais et al. 1981; Marx and Hatchell 1986; Navratil et 

al. 1981; Parker et al. 1986; Riffle and Tinus 1982; Ruehle 1981, 

1982; Ruehle et al. 1981b; Valdes 1986).

A significant increase in survival, stem caliper, or stem height can 

justify the expense of inoculation. Outplanting response to 

inoculation will differ for various habitat types as well as seedling 

host and fungal species (Dixon 1986). On sites that are extremely 

difficult to regenerate (ones that have been planted numerous 

times without seedling survival), seedling survival is of paramount

importance (fig. 5.2.40 and 5.2.41). A successful nursery 

inoculation program begins with the careful evaluation of the need 

for inoculation by the forester, and his/her linking with an 

experienced nursery manager and mycorrhizal specialist to 

produce appropriately inoculated seedlings (Kidd 1982).
Studies in other regions have shown P. tinctorius to be of no 

benefit (Alvarez and Trappe 1983a, Grossnickle and Reid 1982, 

Pilz and Znerold 1986, Ruehle 1983). Clearly no one fungus will 

work well in all situations.

Other fungi have also increased field performance of various 

conifer seedlings, including Cenococcum geophilum (Kropp et al. 

1985, Riffle and Tinus 1982), Laccaria laccata (Thomas and 

Jackson 1983), Suillus bovinus (Ekwebelam and Odeyinde 1985), 

Suillus luteus (Ekwebelam and Odeyinde 1985), Rhizopogon 

luteolus (Ekwebelam and Odeyinde 1985), Rhizopogon roseolus

(Riffle and Tinus 1982), Rhizopogon vinicolor (Castellano and 

Trappe 1985), and Thelephora terrestris (Riffle and Tinus 1982, 

Thomas and Jackson 1983).

Some inoculated fungi do not persist on seedling roots after 

outplanting and thus do not impart any advantage as originally 

designed. For example, in some habitats, Pisolithus tinctorius 

ectomycorrhizae are aggressively displaced from the feeder roots 

of inoculated seedlings by native mycorrhizal fungi after 

outplanting (Dixon et al. 1981, Dixon et al. 1984b, Hung and 

Trappe 1987, McAfee and Fortin 1986, Ruehle 1983). In the 

Pacific Northwest, researchers commonly observe the displacement of Pisolithus tinctorius and other inoculated fungi (Thelephora 

terrestris, Laccaria laccata, and Hebeloma crustuliniforme) after 

outplanting (fig. 5.2.42), most commonly by a Rhizopogon-type 

(Bledsoe et al. 1982, Castellano and Trappe 1987). However, 

some fungi have been shown to persist for several years on inoculated seedlings (Castellano and Trappe 1985, Danielson 1985). 

Persistence of the inoculated mycorrhizal fungus is an important 

criterion when selecting mycorrhizal fungi for inoculation.

129
To prepare the spore inoculum, freshly collected fruiting bodies are 

rinsed with tap water to remove adhering soil or organic matter, 

then cut into pieces (1 to 3 cm3) and blended with tap water at high 

speed for 2 to 3 minutes, until all pieces are thoroughly blended 

(fig. 5.2.48 and 5.2.49). The final consistency is similar to thick 

chocolate milk (fig. 5.2.50). We have found it unnecessary to purify 

spore suspensions. Li and Castellano (1987) and Li (1987) have 

found beneficial microorganisms within and on the surface of 

mature fruiting bodies of various ectomycorrhizal fungi; these 

organisms should be encouraged, not excluded (Garbaye and 

Bowen 1987, Linderman 1988, Schroth and Weinhold 1986).

Spore concentrations within the resulting suspension are

determined with a hemacytometer (blood cell counter) and stored 

in the dark under refrigeration (up to 5 °C or 41 °F) until used. We 

recommend using fresh spores whenever possible, but have 

stored spore suspensions of various Rhizopogon species up to 3 

years without a significant reduction in inoculum effectiveness 

(Castellano 1987).

130

5.2.6 Sources of Inoculum and Inoculation

Techniques

Soil, spores, and vegetative mycelium are the three primary 

sources of ectomycorrhizal and VA mycorrhizal inoculum for 

container seedlings. Each has advantages and disadvantages in 

relation to the objectives and economics of the inoculation 

program. We will discuss ectomycorrhizal fungus inoculum first and 

then VA mycorrhizal fungus inoculum.

5.2.6.1 Soil inoculum

Historically, soil inocula taken from beneath ectomy corrhizal host 

trees have been used extensively, especially in developing 

countries (Mikola 1970). In bareroot nurseries, up to 10% by 

volume of soil inoculum is incorporated into the soil (top 10 cm of 

beds) before sowing (fig. 5.2.43). Goodwin (1976) used 2 ounces 

of screened pine straw as inoculum for loblolly pine container 

seedlings and found a significant increase in height growth after 3 

years in North Carolina. Parke et al. (1983b) reported enhanced 

growth of Douglas-fir container seedlings inoculated with litter and 

humus taken from beneath Douglas-fir trees. This method requires 

large quantities of soil on an annual basis. One of the most serious 

disadvantages of soil inoculum is that weed seeds, rhizomes, and 

potential pathogens may also be inadvertently transported into the 

nursery with the soil. Another disadvantage is the inconsistency of 

the inoculum quality due to varying times and sources of soil 

collection. We do not recommend this method unless other forms 

of inoculum are not available.

5.2.6.2 Spore inoculum

Spores or macerated fruiting bodies of some ectomycorrhizal 

mushrooms, puffballs, or truffles (and false truffles) provide good 

inoculum. Truffles (Ascomycotina) and false truffles 

(Basidiomycotina), from now on together referred to as truffles, are 

uniquely suited for this because their fruiting body tissue consists 

mostly of spore-bearing tissue (fig. 5.2.44-5.2.46), and the fruiting 

bodies can be quite large (fig. 5.2.47).
131
132
133

Spores are applied 6 to 12 weeks after sowing, either with a 

standard watering can (fig. 5.2.51) or through the existing irrigation 

system (fig. 5.2.52). Most truffle spores are less than 50 μm in 

diameter and will pass freely through most filters and nozzle tips. 

The desired amount of spores is mixed into a watering can 

containing sufficient water to cover a certain number or area of 

seedlings (Styroblocks® or racks of plastic tubes). Applying spores 

twice, 2 to 3 weeks apart, works best to assure even distribution 

(fig. 5.2.53), especially when using the irrigation system instead of 

watering cans.

Alternatively, spores can be applied to the seed before sowing 

(Marx and Bell 1985, Marx et al. 1984, Theodorou 1984, 

Theodorou and Benson 1983, Theodorou and Bowen 1973). 

Although we have not tried this method, it may prove more 

effective than the wateringcan method in inoculating each 

seedling. Seed treatment would also allow finer control in matching 

ecotypes of fungi to specific seed sources.
In our recent operational trials, Castellano (1987) has successfully 

inoculated seven million Douglas-fir container seedlings each of the 

last 2 years by adding a spore suspension of Rhizopogon vinicolor 

into the fertilizer injector system. Using the overhead irrigation 

system, a known quantity of spores was applied to blocks of 

250,000 8-week-old seedlings in 5 minutes or less. The treatment 

consisted of a 1-minute prewetting of the growing media, a 

2-minute spore application, and an additional 2-minute wetting of 

the growing media to leach the spores downward into each cavity. 

The additional 2-minute wetting period also serves to rinse the 

water lines in case other fungal isolates or species are to be used 

for different stock.

We have tested many different fungal species using the spore 

suspension method for inoculation; species in the genus 

Rhizopogon succeed the best (table 5.2.5). For (Douglas-fir we 

have focused on Rhizopogon vinicolor, which is host-specific to 

Douglas-fir and has been successfully inoculated as basidiospores 

onto seedlings grown in both bareroot and container nurseries 

(Parke et al. 1983b, Castellano and Trappe 1985, Castellano et al. 

1985). This fungus-host combination produces mycorrhizae with 

prolific rhizomorphs that likely function in water transport 

(Duddridge et al. 1980). Parke et al. (1983a) demonstrated that 

Douglas-fir seedlings inoculated with R. vinicolor withstood and 

recovered from drought better than noninoculated seedlings or 

those inoculated with Pisolithus tinctorius, Laccaria laccata, or an 

unidentified native forest fungus. Most importantly, Douglas-fir 

seedlings inoculated with R. vinicolor survived and grew 

significantly better than noninoculated nursery run seedlings (with 

abundant Thelephora ectomycorrhizae) on routine sites (Castel-

lano and Trappe 1985) and difficult reforestation sites (Castellano 

unpublished data) in southwestern Oregon.

In the southern hemisphere, spores of another Rhizopogon 

species, R. luteolus, have been successfully used to inoculate and 

stimulate growth of pines in Australia (Theodorou 1971; Theodorou 

and Bowen 1970, 1973) and South Africa (Donald 1975).

134
Vegetative inoculum of some fungal species is commercially 

available from Mycorr Tech Inc. (440 William Pitt Way, Pittsburgh, 

PA 15238). Their product comes in 7- to 10-liter bags (fig. 5.2.55), 

is effective (Hung and Molina 1986a, 1986b; Hung and Trappe 

1987); and has a reasonable shelf life (Hung and Molina 1986a). 

Currently, Pisolithus tinctorius, Hebeloma crustuhniforme, and 

Laccaria laccata are readily available (Maul 1985); other 

ectomycorrhizal fungi may be produced as demand warrants.

* The sources of mycorrhizal inocula listed in this chapter are currently 

(1988) the only suppliers of mycorrhizal inocula known to exist. It is not the 

intention of the U.S. Department of Agriculture, or the Forest Service, to 

recommend the products of these companies over any others that may be 

developed in the future.

135

Marx (1976, 1980) and others (Ruehle 1980b) have had similar 

success with inoculating Pisolithus tinctorius onto assorted pine 

species in the southeastern United States. Pisolithus tinctorius has 

stimulated the growth of oak and pine seedlings both in the nursery 

and upon outplanting, particularly on mine tailing or highly eroded 

sites. Although P. tinctorius occurs in limited habitats in the Pacific 

Northwest, it has not performed well in nursery inoculations or 

outplanting trials (Alvarez and Trappe 1983a, 1983b; Castellano 

and Trappe unpublished data).

Spore suspensions of various fungi are available for commercial 

distribution, especially in the Pacific North west, from Forest 

Mycorrhizal Applications (PO Box 385, Murphy, OR 97533).* They 

recently began collecting and distributing spore suspensions of 

various species of Rhizopogon, Suillus, and other ectomycorrhizal 

fungi.

As with vegetative inoculum, not all fungi can be inoculated 

effectively with this method. This inoculum is not free of other 

organisms, but, in the 7 years of our experience with this type of 

inoculum, we have never encountered any harmful effects to the 

seedlings we have treated. The fruiting bodies used for preparing 

the suspension are only seasonally available and, unlike vegetative 

inoculum, the genetic make-up of the spores will vary from year to 

year and place to place.

5.2.6.3 Mycelial inoculum

Over the past few years, much research has concentrated on the 

production and utilization of pure culture inoculum of selected 

ectomycorrhizal fungi. Molina and Palmer (1982) detail isolation 

and maintenance of ectomycorrhizal pure cultures. Marx and 

Kenney (1982) elaborate on production of ectomycorrhizal 

inoculum. Basically, a pure culture of a particular fungus is 

obtained by isolating fungal material (spore germination or 

vegetative tissue explant) onto special media (fig. 5.2.54), that is 

then grown under aseptic conditions to produce inoculum. The bulk 

inoculum, usually produced in a peat-vermiculite carrier moistened 

with nutrient solution, is mixed into container growing media prior to 

filling containers and sowing seed.
136

In another type of noncommercial mycelial inoculum, fungal 

sclerotia are embedded in a liquid or gel base Boyle et al. 1985, 

Boyle et al. 1987, Danielson et al. 1984b, Grenville et al. 1985, 

LeTacon et al. 1983, Mauperin et al. 1987).

Vegetative inoculation has a higher initial cost and requires more 

labor than the spore inoculation method. As with spore inoculation, 

different fungal species also differ in their effectiveness in 

vegetative inoculation. For Example, Rhizopogon vinicolor grows 

well on artificial media but is not effective in colonizing feeder roots 

when used as a vegetative inoculum (Molina 1980).

Marx et al. (1982) used Pisolithus tinctorius in the first wide-scale 

application of ectomycorrhizal inoculum in container nurseries. 

They compared the effectiveness of vegetative inoculum produced 

in their research laboratory against that of a commercially 

produced inoculum on 10 pine species, Douglas-fir, western 

hemlock, and bur oak. Both inoculum sources were effective. 

Inoculum was most effective after leaching with water to remove 

excess nutrients. No other inoculum characteristic or treatment 

significantly affected inoculation success, except that a captan 

drench after seeding improved the effectiveness of the inoculum.

In the Pacific Northwest, other promising fungi such as Hebeloma 

crustuhniforme and Laccaria laccata are easily isolated, grow well 

in pure culture, and when developed on a peat-vermiculite carrier are effective inocula for Douglas-fir container seedlings. Low levels 

of vegetative inoculum are effective even under normal operational 

conditions of abundant water and soluble fertilizer (Hung and 

Molina 1986b). In addition, vegeta tive inoculum of both fungi has a 

shelf life of up to 6 months (Hung and Molina 1986a).

Unfortunately, we have not demonstrated survival or growth 

enhancement, either in the nursery or in planta tions, to justify the 

cost of vegetative inoculum of either H. crustuliniforme or L. 

laccata (Molina 1982, Castellano 1987). Under nursery (Hung and 

Trappe 1987) and field conditions (Castellano 1987) in the Pacific 

Northwest, Laccaria laccata and to a lesser degree Hebeloma 

crustuliniforme ectomycorrhizae are quickly replaced after 

transplanting by indigenous fungi (what appears to be a 

Rhizopogon-type).
VA mycorrhizal fungus pot cultured inoculum is usually added to 

the growing media in one of two methods (see Menge and Timmer 

1982 for additional information). In the first method, the inoculum is 

mixed evenly throughout the growing media prior to filling the 

cavities. In the second method, the inoculum is banded about 3 to 

5 cm (1 .5 to 2 inches) below the surface of the growing medium. 

Although the banding method may be labor intensive, it assures 

rapid contact between the roots and fungus as the roots grow 

down through the inoculum band. Information is variable as to how 

much inoculum is needed to ensure successful inoculation, but, 

from our experience, inoculating with approximately 200 to 500 

spores per seedling is a good beginning for testing an inoculum's 

effectiveness in the nursery. For example Kough et al. (1985) used 

20 ml (0.7 ounces) of pot cultured inoculum (spores + soil + 

chopped roots) to successfully inoculate cedar and redwood 

seedlings growing in 160- cm3 containers; the 20 ml of inoculum 

contained 400 to 770 spores and 30 to 68% of the root pieces 

were colonized. VA mycorrhizal fungi are sensitive to high levels of 

fertilizer, as are many ectomycorrhizal fungi. Thus, careful 

monitoring of mycorrhizal development under various management 

practices will be needed to develop compatible regimes.

137

5.2.6.4 Vesicular-arbuscular inoculum

Two major features of VA mycorrhizal fungi greatly influence both 

natural and artificial means of seedling inoculation. First, as noted 

in section 5.2.1.2, VA mycorrhizal fungus spores are not typically 

wind dispersed like many ectomycorrhizal fungus spores; VA 

mycorrhizal fungus spores will not blow in from outside the nursery, 

or from within the nursery, to natu rally inoculate seedling. Thus, VA 

mycorrhizal host plants grown in artificial growing media or 

sterilized soil will not form mycorrhizae. Second, since VA 

mycorrhizal fungi cannot be grown in pure culture (that is, in 

absence of a host), bulk vegetative (mycelial) inoculum, as 

currently produced for ectomycorrhizal fungi, is not available. 

Nonetheless, other inoculum sources and techniques are available 

for VA mycorrhizal fungi and in many ways parallel those used for 

ectomycorrhizal fungi.

Taking soil from beneath VA mycorrhizal hosts in nature and 

incorporating it into the container substrate is a simple inoculation 

method. Parke et al. (1983b) reported enhanced growth of western 

redcedar container seedlings inoculated with litter and humus taken 

from beneath Douglas-fir trees. However, as for ectomycorrhizal 

inoculation, we discourage this technique because of the potential 

of introducing unwanted pests into the nursery and the large 

quantity of soil needed.

Although we cannot, as yet, produce pure vegetative cultures of VA 

mycorrhizal fungi, we can still mass produce fungus inoculum by 

allowing a known VA fungus to grow in association with a host and 

then use the soil and roots as inoculum. This procedure is called 

"pot culturing." In general, spores of a particular VA mycorrhizal 

fungus are first retrieved from natural soil by various separation 

techniques (see Ferguson and Woodhead 1982), identified, surface 

sterilized, and then added into a sterile growing medium in which a 

host like sorghum or clover is then grown. As the plant grows, it 

forms VA mycorrhizae with the desired fungus; the fungus then 

spreads through the growing medium and produces abundant 

spores. These spores can then be retrieved from the growing 

medium for use as inoculum (fig. 5.2.56), or, more commonly, the 

entire growing medium with the mycelium, spores, and roots 

(chopped) it contains can be used as inoculum.
Pot cultured inoculum provides the best source of VA mycorrhizal 

fungi for several reasons. If the pot cultured inoculum is grown 

properly, there is little risk of introducing unwanted pests or 

pathogens. The inoculum is usually reliable, efficient, and easily 

introduced into growing media. Most importantly, pot culturing 

allows the use of selected highly beneficial fungus strains to 

provide maximum enhancement of seedling growth and survival 

(fig. 5.2.57 and 5.2.58). Considerable research has been 

conducted and is currently in progress on selection of beneficial VA 

mycorrhizal fungi for plant inoculation. Although the majority of this 

research has been done with agricultural crops, information is also 

available for VA mycorrhizal forest tree species (see Brown et al. 

1981; Kormanik 1985; Kormanik et al. 1977, 1981, 1982; and 

Kough et al. 1985).

A commercial source of VA mycorrhizal fungi is now available; 

others continue to be developed. One promising source of 

inoculum is being developed and marketed by NPI (417 Wakara 

Way, Salt Lake City, Utah 84108). They are able to produce 

inoculum of several VA mycorrhizal fungi (fig. 5.2.59) and are 

developing methods for bulk production of axenically grown 

inoculum free of pathogens (Wood 1987). NPI is also involved in 

site reclamation, so their experience with incorporating microbial 

inoculants into plant rearing programs will be an added source of 

consultation for nurseries wanting to begin VA mycorrhizal 

inoculation programs.

138
As with implementing an ectomycorrhizal inoculation program, 

nursery managers should have clear objectives for VA mycorrhizal 

inoculations. VA mycorrhizal inoculation can improve growth in the 

nursery and reduce fertilizer costs; inoculated stock can also 

perform better than noninoculated stock, especially when planted 

into environmentally stressful habitats or where native VA 

mycorrhizal fungi are lacking (Johnson 1987). Whatever your 

objectives, working with knowledgeable specialists to aid in 

selection of VA mycorrhizal fungi, techniques of inoculation, and 

evaluation of inoculation success is strongly recommended.

139

5.2.6.5 Isolate selection and ecotypic variation

Tables 5.2.6 (by fungus) and 5.2.7 (by host) list the many different 

fungus--host combinations that have been successfully inoculated 

onto container seedlings. The response of the seedling host can 

vary considerably. Of the 118 successful fungus-host combinations 

listed, only 105 combinations have growth characteristics reported 

for comparison. Over one-third of the fungus--host combinations 

stimulated seedling growth, whereas nearly a fourth reduced 

seedling growth. Six percent increased and decreased growth of 

the same seedling host in different trials. For the most part, growth 

of hardwoods (especially oaks) was consistently stimulated by 

fungal inoculation, whereas growth of pines, spruces, firs, and 

Douglas-fir seedlings was more often not affected or suppressed 

rather than stimulated. Growth of larch seedlings was unaffected 

by inoculation. Hebeloma crustuliniforme, and Laccaria laccata

reduced seedling growth more often than it increased it. Pisolithus 

tinctorius stimulated a majority of the responsive hosts. Although 

many of these symbionts had little or no effect on container 

seedling growth in the nursery, some of these symbionts 

stimulated increased seedling field performance (Thomas and 

Jackson 1983). The nursery manager with advice from a 

mycorrhizal specialist can select fungus-host combinations that 

have promise to meet objectives for a particular host species.

Mycorrhizal fungi constantly compete with other mycorrhizal fungi 

and microorganisms for living space in the seedling rhizosphere. 

just as some mycorrhizal fungi can antagonize pathogens, so can 

some mycorrhizal fungi antagonize other mycorrhizal fungi. In pure 

culture some Rhizopogon species produce chemicals that inhibit

such fungi as Cenococcum geophilum, Hebeloma crustuliniforme, 

Laccaria laccata, Pisolithus tinctorius, and Thelephora terrestris 

(Castellano 1987). Understanding competitive interactions between 

mycorrhizal fungi will allow us to select fungal species or isolates 

for their ability to dominate root systems upon inoculation and 

continue to provide selected benefits to the inoculated seedlings 

when outplanted.
140
141
142
143
144
145
146
147
148

The influence of fungal genetic composition on the ability of a 

fungal species to form mycorrhizae with hosts from different seed 

sources has not been studied. Even within a fungal species, 

isolates from different habitats have different morphological 

characters (fig. 5.2.60 and 5.2.61). The applicability of inoculating 

a specific seed source of seedling host with an ecotype of a 

particular fungus has the potential of matching fungi and seedling 

host to habitat. Different genotypes of Scotch pine (Lundeberg 

1968), lodgepole and ponderosa pine (Cline and Reid 1982), 

Sitka spruce Walker et al. 1986), European larch (Zhu and 

Navratil 1987), and Douglas-fir (Wright and Ching 1962) formed 

significantly differing amounts of ectomycorrhizae when 

inoculated with the same fungal isolate and grown under common 

conditions. Growth response of the seedling host can also differ 

(Cline and Reid 1982, Zhu and Navratil 1987). Pisolithus tinctorius

(Dixon et al. 1984a, Marx 1981, Molina 1979), Suillus granulatus

(Dixon et al. 1984a) and Hebeloma crustuliniforme (Molina 1987) 

exhibit the same varying pattern of response (host growth or 

ectomycorrhizal formation) when the same seed source but 

different fungal isolates are used for inoculation, but Laccaria 

laccata does not (Molina 1982). The mycorrhizal fungus and tree 

host have co-evolved to some degree within their geographic cal 

(ecotypic) realm. Mycorrhizal research programs are currently 

investigating the importance of matching ecologic adaptations of 

trees and fungi for wide-scale application.
5.2.7 Evaluating Inoculation Success

Given the diversity in crop species, growing conditions, and 

management techniques in container nurseries, mycorrhizal 

inoculations that work well in one nursery may not work well in 

others. We urge each nursery to test recommendations for 

mycorrhizal management in their nurseries on a small scale 

before trying to inoculate the entire nursery. A few thousand 

seedlings are more than enough for first inoculation attempts. Be 

sure to incorporate some variation from the standard inoculation 

procedure into test programs, for example, vary spore rates, timing 

of application, fertilizer levels, and types of fertilizer so that you can 

learn more about how your management practices interact with 

inoculation success. Also, try to work with a scientist or statistician 

who can help develop a simple experimental design to facilitate the 

analysis of results.

5.2.7.1 Rating mycorrhizal formation

Nursery managers or growers should ideally have some basic 

training in identification of mycorrhizal types. This experience can 

be through one-on-one training with an expert or at a workshop. 

Even after training, nursery managers and growers should send 

some of their treated seedlings to a recognized expert in the field 

to corroborate their findings. Helpful hints are supplied in section 

5.2.3.

5.2.7.2 Designing outplanting trials

The true test of a nursery inoculation program may not be 

apparent until after the seedlings are outplanted. Because 

seedling size is a major factor in outplanting success, measure 

seedlings before outplanting to assure that you are comparing 

seedlings of similar size in the field (fig. 5.2.62). In one study 

(Barnett 1982), differences in initial seedling size were more 

closely related to field performance than amount of Pisolithus tinctorius ectomycorrhiza. Also realize that rough handling of 

seedlings during any phase of the planting process can be 

detrimental to ectomycorrhizae (Tabbush 1986).

149
5.2.7.3 Economic considerations

Cost effectiveness is diffi cult to generalize because it depends on 

type of inoculation, individual nursery management costs, and 

scale of inoculation. Most importantly, these calculations are 

compounded by the specific inoculation objectives and definition 

of effectiveness. For example, a nursery trying to increase stern 

caliper and seedling uniformity or reduce culls will judge 

effectiveness (and its costs) differently than a nursery whose 

objective is to improve seedling field performance. The former 

deals with immediate benefit and costs while the latter deals with 

an effectiveness concept tied to the future. Nursery managers 

need to calculate specifics of cost-effectiveness as they develop 

individual inoculation programs. This is another reason to keep the 

scale of first inoculations small.

A new company is now preparing spore suspensions of various 

fungi for commercial distribution, especially in the Pacific 

Northwest. Forest Mycorrhizal Applications (1032 Starlite, Grants 

Pass, OR 97526) has recently begun collecting and distributing 

spore suspensions of various species of Rhizopogon, Suillus, 

and other ectomycorrhizal fungi. The 1988 cost of the inoculum 

was from 0.25 to 0.95¢ per thousand seedlings; application is 

additional (see section 5.2.6.2).

Mycorr Tech Inc. (440 William Pitt Way, Pittsburgh, P~ 15238) is 

presently supplying commercial vegetative inoculum. In 1988, 

their product cost approximately $1.00 to 2.00 per thousand 

seedlings; application is additional. Tests have shown their 

product to be reliable, reproducible, relatively quickly available, 

and uncontaminated (see section 5.2.6.3).

Commercial sources of VA mycorrhizal fungi are now available 

and continue to be developed. NPI (417 Wakara Way, Salt Lake 

City, Utah 84108) produces inoculum of several VA mycorrhizal 

fungi. In 1988, their product costs approximately $2.00 to 5.00 

per thousand seedlings, depending on inoculation procedure. We 

have noted that product costs have steadily decreased during 

the last 2 years (see section 5.2.6.4).

The outplanting tr ial design should be simple and straightforward. A 

randomized block design with 3 to 5 blocks spread across a 

representative area of a specific habitat type will generate enough 

information to extrapolate to similar sites. The blocks must be small 

enough to reduce within-block variation due to micro- site but large 

enough to provide meaningful replication. When in doubt consult 

with a statistician. We find that 20 to 50 seedlings of each treatment 

combination (noninoculated or inoculated) per block are usually 

adequate. Blocks should be separated from one another with 3- to 

6-m (10- to 20-foot) buffer strips. Spacing of seedlings can be 

critical. We have used spacing as close as 1.2 x 1.2 m (4 x 4 feet) 

but prefer 2.4 x 2.4 or 3.6 x 3.6 m (8 x 8 or 12 x 12 feet) to correlate 

with what is done operationally. Within the blocks, seedlings from 

the same treatment are planted in rows of 10 to 25, with row 

location randomized within each block. It is helpful in subsequent 

years to have the block corners marked with 1.2-m (4-foot) metal or 

plastic stakes (not wood, which breaks easily) and to mark the 

beginning and end of each seedling row with heavy-gauge wire 

stakes. Treatment codes on metal tags can be attached to the 

stakes at the beginning of each row. Protecting the seedling is 

critical (fig 5.2.63): many vigorous inoculated seedlings are lost to 

deer browsing because they are more palatable than noninoculated 

control seedlings. Measurements of seedling height and stem 

caliper at time of outplanting provide baseline data to calculate 

future annual increment of growth. Measurements are taken 

anytime during seedling dormancy depending upon site 

accessibility.

We find that at the beginning of the second year we have a more 

accurate comparison of inoculation treatments than the first year 

because first year seedling growth will be influenced by nursery 

practices (that is, fertilization and watering regimens) of the 

previous year. Typically, measurements are taken for the first 5 

years; evaluation of these data determines if monitoring should 

continue beyond that. Although not routine, we encourage 

excavation of seedling root systems to allow observation of the 

persistence of inoculated fungi on old feeder roots and their growth 

onto new feeder roots. Five to ten seedlings per inoculation 

treatment per year is adequate. Techniques for examining mycorrhizae on root systems from the field are similar to those discussed 

in section 5.2.3.

150
151
The effects of CuCO3 and IBA on ectomycorrhizal fungus 

inoculation have been determined for ponderosa, lodgepole 

(McDonald et al. 1981), loblolly, longleaf, shortleaf, and eastern 

white pine (Ruehle 1985a). In all cases, 50 g of CuCO3 /liter of 

latex paint was used. Treated ponderosa and lodgepole pine seedlings inoculated with Suillus granulatus or Pisolithus tinctorius had 

somewhat larger stem height and caliper and significantly reduced 

root deflections compared to nontreated seedlings. CuCO3

treatment of remaining pine species had little effect on seedling 

growth, except that feeder root formation was usually stimulated. 

Formation of ectomycorrhizae was either not affected (loblolly and 

shortleaf pine), stimulated (longleaf pine), or depressed (eastern 

white pine) (Ruehle 1985a). In a follow-up outplanting trial, P. 

tinctorius-inoculated loblolly and longleaf pine grown in containers 

treated with CuCO3 survived and grew better than untreated P. 

tinctorius-inoculated seedlings on a routine reforestation site in the 

southeast United States (Ruehle 1987).

Copper sulfide has also been used to prevent root spiraling in 

Chinese pine seedlings grown in polyethylene-coated kraft paper 

containers (Dong and Burdett 1986). Unfortunately the effects of 

the chemical on ectomycorrhizal inoculation were not explored.

Nursery managers may want to try some of these feeder root 

enhancement techniques on a small scale and carefully monitor 

the effects on root growth and mycorrhizal development before 

wide-scale application.

5.2.8.2 Fertilizer

Mycorrhizae and mycorrhizal fungi are extensions of a plant's root 

system; they extract nutrients and water from soil and translocate 

them to the host. Plants respond to mycorrhizal formation most 

strongly in soils of low fertility. It follows that most mycorrhizal fungi 

are adapted to the infertile conditions of forest soils. Many 

mycorrhizal fungi do not grow well in artificial growing media that 

are frequently drenched with high levels of soluble fertilizer or 

amended with slow-release fertilizer. Inhibition of mycorrhizae by 

high levels of fertilization plus the lack of mycorrhizal fungus 

propagules in artificial growing media pose the greatest challenge 

to mycorrhiza management programs.

152

5.2.8 Factors Affecting Mycorrhizal

Development

5.2.8.1 Root development

The major lateral roots of conifers grown in containers typically 

grow out to the container wall and then downward parallel to the 

container side for their first 10 to 15 cm. This growth form 

discourages initiation of secondary laterals; many downward 

trained roots continue in this fashion after outplanting. On the 

outplanting site, the upper portion [10 to 15 cm (4 to 6 inches)] of 

the soil profile usually has high oxygen, moisture, and nutrient 

availability and thus is conducive to high rates of microbiological 

activity (Harvey et al. 1987). To insure seedling establishment after 

outplanting, exploration of the upper soil layers by feeder roots and 

ectomycorrhizae is desirable.

Nursery techniques to manipulate root form of container seedlings 

and enhance root growth potential of seedlings after outplanting 

are relatively new. One involves coating the inside of the container 

with a root pruning chemical contained within a latex paint. After 

the paint dries the containers are filled with growing media and 

sown in normal fashion (Romero et al. 1986). Various 

concentrations of three different chemi cals have been tried. 

Trifluralin (a herbicide) at all concentrations tested (0.56 to 70.88 

g/I of paint) adversely affected ponderosa pine seedlings 

(McDonald et al. 1981). A 5-g/I concentration of indolebutyric acid 

(IBA) applied to the container wall increased ponderosa pine 

seedling growth somewhat, but growth was weak and erratic 

compared to container wall treatment with 50 g/I concentration of 

cupric carbonate (CuCO3) (McDonald et al. 1984). Seedlings 

grown in containers treated with CuCO3 and then transplanted and 

grown for an additional 5 weeks had 27% of their new roots as 

side roots, while the untreated seedlings produced only 8%. 

Seedlings treated with 100 g/I of CuCO3 had significantly higher 

shoot and root dry weight and larger stem height than seedlings 

treated with 0.1 g/I, and also had one fourth (3.7 vs. 12.2) as many 

roots deflected down the container wall (McDonald et al. 1981). 

Unfortunately, some latex paint carriers can be phytotoxic, with the 

detrimental effects overcome only at the higher CuCO3

concentrations. Other potential carriers need to be tested.
153

Fertilizer form is also important; compared to nitrate-N, 

ammonium-N is usually better utilized by a variety of mycorrhizal 

fungi (Bledsoe and Zasoski 1983, Littke et al. 1984, Harley and 

Smith 1983). Ammonium-N fertilization decreases the pH of the 

growing media whereas nitrate-N fertilization will increase the pH 

of the growing media. As we will see later, many ectomycorrhizal 

fungi prefer acidic growing conditions, so fertilization with nitrate-N 

will adversely affect inoculation of alkaline-sensitive fungi.

Given variable responses to fertilizers by different mycorrhizal 

fungi, we cannot recommend specific levels, types, or forms of 

fertilization to promote mycorrhizal development on container 

seedlings. Optimum fertilization levels must be determined by 

each nursery manager, depending on whether the objective is 

promoting mycorrhizal development of naturally occurring fungi or 

ensuring inoculation with a particular fungus. Nursery managers 

should also realize that mycorrhizal fungi may provide seedling 

growth stimu lus equal or similar to high levels of fertilization and 

thus result in a fertilizer cost saving. If enhancing outplanting 

performance via mycorrhizal inoculation is a goal, close control 

over the fertility and how it is applied is essential.

Mycorrhizal management should be considered as part of the 

overall container tree seedling culture. Be open minded about 

modifying fertilization levels, application schedules, and fertilizer 

forms to meet mycorrhizal management objectives. Nursery 

managers and staff are highly skilled in developing the optimum 

cultural practices to produce vigorous planting stock; encouraging 

mycorrhizal development on container stock requires these same 

skills.

Because various species of mycorrhizal fungi respond differently to 

fertilization, fungi adapted to nursery fertility conditions can be 

used or fertilizer application can be modified to promote 

colonization by a desired, but fertilizer-sensitive fungus. For 

example, high levels of soluble NPK fertilizer reduce 

ectomycorrhizal forma tion by Pisolithus tinctorius (Crowley et al. 

1986, Danielson et al. 1984a, Dixon et al. 1985, Ekwebelam and 

Reid 1983, Maronek et al. 1981, Maronek et al. 1982, Marx et al. 

1982, Pope and Chaney 1984, Ruehle 1980a, Ruehle and Wells 

1984, Rupp and Mudge 1985). Reducing fertility levels by half may 

double ectomycorrhizal colonization for some hosts (see Marx et 

al. 1982). On the other hand, some fungi such as Laccaria laccata 

and Rhizopogon vinicolor are little affected by high levels of 

soluble fertilizer. Inoculation with these fungi in commercial 

nurseries has been successful without altering the routine 

fertilization regime (Castellano et al. 1985, Danielson et al. 1984a, 

Hung and Molina 1986a, Molina and Chamard 1983, Tyminska et 

al. 1986).

Vesicular-arbuscular mycorrhizal formation of container 

yellow-poplar (Verkade and Hamilton 1985) and south ern 

magnolia (Maronek and Hendrix 1978) seedlings has been 

encouraged by certain fertilization regimes.

Fertilizer type can also affect mycorrhiza development. The two 

common types of fertilizer, soluble and slowrelease, have been 

shown to affect the successful outcome of ectomycorrhizal 

inoculation (Castellano et al. 1985, Maronek et al. 1982). 

Castellano et al. (1985) found that the inoculation success of 

Rhizopogon vinicolor spore application on Douglas-fir container 

seedlings was reduced by slow-release fertilizer but not by soluble 

fertilizer. As recommended in volume four of this series, we advise 

against the use of slow-release fertilizer due to the unknown 

aspect of what the seedlings are actually exposed to by way of 

fertilizer nutrients.

Although foliar application of NPK is not routinely used in container 

nurseries, black oak seedlings receiving foliar NPK had 

significantly greater Pisolithus tinctorius ectomycorrhizae and 

fructose content of feeder roots compared to the soluble NPK 

drench treatment (Dixon et al. 1981).
To avoid water roots, and thus encourage good development of 

feeder roots and ectomycorrhizae, nursery managers must 

regularly examine root systems and modify watering regimes as 

appropriate. As emphasized before, this must become a regular 

practice when assessing root and overall seedling quality 

throughout the growing season.

5.2.8.3 Water

Either too much or too little water reduces feeder root formation 

(Ruark et al. 1982), especially in Douglas-fir and spruce. Many 

nurseries water their seedlings to growing medium saturation every 

day (Matthews 1983). One symptom of over-irrigation is the 

formation of water roots-thick fleshy, opaque nonmycorrhizal roots 

that lack root-hairs (fig. 5.2.64). These water roots act as giant 

sponges that readily absorb water and soluble nutrients. They lack 

the feeder roots needed for mycorrhizal formation (Castellano 

1987, Dixon et al. 1985) and are essentially nonfunctional in water 

and nutrient uptake upon outplanting (Castellano 1987, Dixon et al. 

1983). Water roots have been observed to die and decompose 

soon after outplanting (G. Hunt 1987). These water roots are 

sometimes seen in extreme situations, usually compacted growing 

media. Heavy irrigation with good porous growing media will not 

cause problems. Peat quality is critical: poor peat with a high 

percentage of "fines" will cause growing media to drain poorly. 

Also, xylem-girdling insects can cause water roots by restricting 

water flow to the shoot. From our experience, some inoculation 

experiments have failed because the fungus did not have the 

opportunity to form ectomycorrhizae due to excessive water roots. 

Root dry weight is not a good indication of root quality; a root 

system with large water roots may have the same dry biomass as 

one with many small feeder roots.

Seedlings that are somewhat overwatered (but not to the point of 

having excessively swollen roots) develop many unbranched or 

poorly branched laterals near the surface of the container walls 

and at the container bottom. In these seedlings, optimum 

development of feeder roots and thus mycorrhizae occurs only in 

the inside portion and near the top of the plug where aeration is 

best. These seedlings have extremely poor root regeneration 

potential upon outplanting.

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155

5.2.8.4 Growing media

The physical and chemical makeup of the growing media will 

influence success of mycorrhizal inoculation programs. Pore size 

and distribution and pH (optima and tolerance) will directly affect 

not only feeder root formation (Ruark et al. 1982) and distribution 

(fig. 5.2.65) but also ectomycorrhizal development. Compacted 

growing media not only inhibit feeder root initiation but also inhibit 

lateral and feeder root extension. The high percentage of peat 

moss in most artificial growing media affects their physical and 

chemical properties, that is, pH. Our field observations infer that 

some ectomycorrhizal fungi prefer soils with high organic matter 

contents (for example, decomposed logs, with pH = 4), whereas 

others grow well in mineral soils with greatly reduced amounts of 

organic matter (for example, recently burned areas, with pH = 7). 

Ectomycorrhizal fungi have various pH optima for growth in pure 

culture as well (Hung and Trappe 1983). Some fungi grow equally 

well over a relatively wide pH range, whereas others are less 

tolerant (Hung and Trappe 1983). For example, Pisolithus tinctorius

formed more ectomycorrhizae at pH 5.5 than at 6.5 when 

inoculated onto pecan seedlings (Sharpe and Marx 1986).

Growing medium compaction does not seem to elimi nate fungal 

growth, but it greatly reduces formation of feeder roots needed for 

ectomycorrhizal colonization. The container growing media should 

provide adequate pore space for oxygen exchange to promote 

vigorous growth by both roots and fungi. We recommend selecting 

fungi that grow well over a wide range of growing media pH for 

nursery inoculation.
5.2.8.5 Temperature

As with pH, ectomycorrhizal fungi have tolerance ranges and 

optima for temperature (Hacskaylo et al. 1965, Marx and Bryan 

1971, Marx et al. 1970, Samson and Fortin 1986). Growing 

medium temperatures in containers can vary widely, from cold [0 

°C (32 °F)] in winter or during preplanting storage to hot [38 °C 

(100 °F)] during summer. Some mycorrhizal fungi will tolerate this 

wide temperature fluctuation during seedling production, others will 

not. For example, aseptic loblolly pine seedlings inoculated with 

Thelephora terrestris or Pisolithus tinctorius grew well and formed 

abundant ectomycorrhizae at 25 °C (77 °F). When these same 

seedlings were transferred to a room with 40 °C (104 °F) soil 

temperatures, the T. terrestris--inoculated seedlings died or 

declined while P. tinctorius-inoculated seedlings thrived (Marx and 

Bryan 1971).

Ectomycorrhizal feeder roots also differ in ability to withstand cold. 

In many nurseries, preplanting cold storage of seedlings is 

common. P. tinctorius ectomycorrhizae survived cold storage on 

shortleaf pine (Marx 1979a) but not on ponderosa pine (Alvarez 

and Linderman 1983) or Douglas-fir (Castellano unpublished data). 

Cold storage of P. tinctorius vegetative inoculum decreases its 

effectiveness, while inoculum of Laccaria laccata and Hebeloma 

crustuliniforme formed abundant ectomycorrhizae after cold 

storage (Hung and Molina 1986a).

Knowledge of the temperature tolerance of various fungi 

must be used in selecting a fungus for your inoculation 

program.

5.2.8.6 Pesticides

Pesticides cause a multitude of complex reactions on target and 

nontarget organisms. Generalizations about reactions to pesticides 

must be approached with caution. For example, pesticides that 

affect mycorrhizal fungi or mycorrhiza development can affect 

seedling growth response either for better or for worse. Trappe et 

al. (1984) review effects of pesticides on mycorrhizal fungi and 

mycorrhiza development. Tables 5.2.8 to 5.2.11 are condensed 

from Trappe et al. (1984) for reference to container nurseries.

Sterilants. Artificial growing media are generally considered to be 

"essentially sterile" and therefore, sterilants are not normally used 

in container nurseries. Because of recent problems with root 

diseases, however, some nursery managers are beginning to 

sterilize their growing media and containers (table 5.2.8). Methyl 

bromide--chloropicrin mixes are effective sterilants, and under 

optimum conditions of application they nearly eliminate both 

beneficial and pathogenic soil organisms from the treated growing 

media. Optimum conditions are rare, however, so soil organisms 

are rarely completely eliminated. Methyl bromide fumigation is 

used in bareroot seedling mycorrhiza inoculation programs to 

reduce competition by wild fungi with the inoculated fungus. For 

the artificial growing media (for example, milled bark) in container 

nurseries, steam pasteurization serves the same purpose 

effectively.

Fungicides. Most fungicides are selective for certain groups of 

fungi (table 5.2.9). The thiazoles (benomyl, carbendazim, and 

fuberidizole) will suppress Zygomycotina but are less detrimental or 

even stimulatory (Pawuk and Barnett 1981, Pawuk et al. 1980) to 

most Basidiomycotina or Ascomycotina. Since VA mycorrhizal 

fungi are Zygomycotina, special attention needs to be paid to the 

application of this group of fungicides in nurseries where VA 

mycorrhizal hosts are grown. The thiazoles would be the fungicides 

of choice for nurseries growing ectomycorrhizal hosts (Pinaceae). 

The dithiocarbamates (ferbam, mancozeb, zineb, and ziram) and 

substituted aromatics tend to inhibit mycorrhizal fungi of both 

groups. The dicarboximides (captafol and captan) are usually not 

inhibitory at low application rates (see table 5.2.8) but can be at 

higher application rates (Pawuk et al. 1980) or can even be 

stimulatory to both groups of mycorrhizal fungi (Owston et al. 

1986).

156
157

The importance of choosing chemicals for pest control carefully is 

illustrated by programs to control fusiform rust on southern pine 

seedlings inoculated with Pisolithus tinctorius. Ferbam has been 

used to control fusiform rust in southern forest nurseries, but it 

requires repeated applications to be effective. Recently, bayleton 

has proven effective in fusiform rust control and is applied only a 

few times during the growing season. Although bayleton costs 

more than ferbam, the fewer applications reduce labor for a 

significant savings over use of ferbam. Unfortunately, bayleton 

selectively inhibits formation of Pisolithus tinctorius ectomycorrhizae compared to naturally occurring ectomycorrhizal fungi (Kelley 

1987, Marx and Cordell 1984, Rowan 1984). Hence it works 

against Pisolithus inoculation success.

Seed treatment with fungicides appears not to affect 

ectomycorrhizal development following germination, unless the 

seeds are coated with ectomycorrhizal fungus spores 

(Theodorou and Skinner 1976). Fungicidal treatment of seeds of 

VA mycorrhizal hosts can negatively affect VA mycorrhizal 

development following germination, however Ualali and Domsch 

1975).
Herbicides. Interpreting results from herbicide trials is difficult 

because effects on the host plant can indirectly affect the 

mycorrhizal fungus. Usually, herbicide concentrations that 

significantly affect mycorrhizal fungi are considerably higher than 

recommended application rates (table 5.2.10).

Some herbicides, like simazine, actually stimulate growth of 

mycorrhizal fungi in axenic culture as well as under field 

conditions.

Insecticides and nematicides. Generally, high concentrations of 

insecticides or nematicides inhibit fungal growth in pure culture 

(table 5.2.11). Relatively little information is available on effects of 

these compounds on mycorrhizal fungi, however, so we cannot 

provide firm recommendations.

General pesticide recommendations. The literature on 

interaction of pesticides with mycorrhizal fungi and subsequent 

mycorrhizal development is confusing and incomplete. Much work 

is needed to understand why one host-fungus combination is 

affected in certain conditions and another is not. Careful 

observation and recordkeeping by the nursery manager is 

important for integrating mycorrhizal management into the total 

nursery operation. Growers must ascertain what and how much 

pesticide will affect their various crops under specific growing 

conditions. The literature provides a guide to some of the potential 

incompatibilities between pesticide, substrate, host, environment, 

and mycorrhizal fungus.

158
159

• Regularly examine and keep careful records of feeder 

root and mycorrhizal development of different stock 

throughout the nursery. Correlate this information with 

records of other nursery practices to become familiar 

with how one influences the other.

• Explore the various options for inoculation that are 

available when the need for an inoculation program 

develops, and seek the advice of a mycorrhizal specialist 

for actual implementation.

• Experiment wisely with inoculations, beginning on a small 

scale and with well-designed studies that include 

controls.

• Keep abreast of current progress in mycorrhizal 

technology through reading, attending workshops, or 

consult with a mycorrhizal specialist periodically.

• Obtain the reference text, Methods and Principles of 

Mycorrhiza Research, published by the American 

Phytopathological Society (Schenck 1982).

• Include some measure of mycorrhizal development in 

assessing the overall quality of your seedlings.

• Finally, let your customers know about your inoculation 

program and its benefits, because good mycorrhizal 

development is an additional selling point to the 

commercial market.

5.2.9 Conclusions and Recommendations

We cannot overemphasize that mycorrhizae must be included in 

any assessment of root development and seedling quality. Trees 

have co-evolved with and become dependent upon their 

mycorrhizal associations for survival and healthy growth in all 

forestry settings. Foresters and nursery managers are well aware 

of the critical stress period that seedlings experience at transplanting. Thus, it is of the utmost priority that nurseries grow and 

send to the reforestation site seedlings with abundant mycorrhizae 

on their root systems. Seedlings without mycorrhizae will have to 

form them before the seedlings can begin to actively take up water 

and nutrients from the soil. Thus, seedlings with mycorrhizae are 

better prepared to immediately begin soil exploration and so stand 

a better chance for survival and early growth than nonmycorrhizal 

seedlings.

Considerable research on mycorrhizal applications in forestry is 

now in progress. A primary focus continues to be the selection of 

fungi for nursery inoculation based on specific ecological benefits, 

for example, providing drought tolerance. Another research 

direction concentrates on how much natural fungus inoculum is left 

on variously disturbed reforestation sites. This latter direction is 

extremely important because it will help foresters predict which 

reforestation sites may be suffering from a natural mycorrhizal 

fungus deficiency and thus need inoculated nursery stock. In the 

future, both research directions will provide nursery and forest 

management tools to enhance tree regeneration programs 

worldwide.

To reach these goals we offer the following recommendations to 

aid nursery managers in incorporating mycorrhiza management 

practices into their overall seedling production programs. The 

nursery staff should:

• As a first step, learn the basic biology of mycorrhizae, 

understand why they are important, and be aware of the major 

benefits they provide plants.

• Learn to recognize mycorrhizae, identify different types, and 

quantify the amount of mycorrhizae on a seedling root system.

• Understand that nursery practices, especially watering, 

fertilization, and pesticide application, affect mycorrhizal 

development in order to avoid negative impacts.
Berry, C.R. 1982. Survival and growth of pine hybrid seedlings 

with Pisolithus ectomycorrhizae on coal spoils in Alabama and 

Tennessee. Journal of Environmental Quality 11: 709-715.

Bledsoe, C.S.; Tennyson, K.; Lopushinsky, W. 1982. Survival 

and growth of Douglas-fir seedlings inoculated with 

mycorrhizal fungi. Canadian Journal of Forest Research 12: 

720-723.

Bledsoe, C.S.; Zasoski, R.J. 1983. Effects of ammonium and 

nitrate on growth and nitrogen uptake by mycorrhizal 

Douglas-fir seedlings. Plant and Soil 71: 445-454.

Bowen, G.D. 1973. Mineral nutrition of ectomycorrhizae. In: Marks, 

G.C.; Kozlowski, T.T. (eds.). Ectomycorrhizae, their ecology 

and physiology. New York: Academic Press: 151-205.

Boyle, C.D.; Gunn, K.L.; Robertson, W.J. 1985. Development of 

methods for the production of mycelial slurry inoculum. In: 

Molina, R. (ed.). 6th North American Conference on 

Mycorrhizae. 1984 June 25-29; Bend, OR. Corvallis, OR: 

Forest Research Laboratory: 225.

Boyle, C.D.; Robertson, W.J.; Brown, H.L. 1987. Mycelial 

suspensions as ectomycorrhizal inoculum. In: Sylvia, D.M.; 

Hung, L.L.; Graham, J.H. (eds.). 7th North American 

Conference on Mycorrhizae. 1987 May 3-8. Gainesville, FL. 

Gainesville, FL: University of Florida: 85.

Branzanti, B.; Zambonelli, A. 1987; Ectomycorrhizal inoculation of 

containerized Pinus pinaster seedlings. In: Sylvia, D.M.; Hung, 

L.L.; Graham, J.H. (eds.). 7th North American Conference on 

Mycorrhizae. 1987 May 3-8. Gainesville, FL. Gainesville, FL: 

University of Florida: 124.

Brown, R.W.; Schultz, R.C.; Kormanik, P.P. 1981. Response of 

vesicular-arbuscular endomycorrhizal sweetgum seedlings to 

three nitrogen fertilizers. Forest Science 27: 413-420.

Burdsall, H.H., Jr.; MacFall, J.S.; Albers, M.A. 1986. Hebeloma 

arenosa (Agaricales, Corti nariaceae), a new species from 

Lake States nurseries. Mycologia 78:862-865.

Castellano, M.A. 1987. Unpublished data.

Castellano, M.A.; Trappe, J.M. 1987. Unpublished data.

Castellano, M.A.; Trappe, J.M. 1985. Ectomycorrhizal formation 

and plantation performance of Douglas-fir nursery stock 

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5.2.10 References

Alvarez, I.F.; Linderman, R.G. 1983. Effects of ethylene and 

fungicide dips during cold storage on root regeneration and 

survival of western conifers and their mycorrhizal fungi. 

Canadian Journal of Forest Research 13: 962-971.

Alvarez, I.F.; Trappe, J.M. 1983a. Dusting roots of Abies 

concolor and other conifers with Pisolithus tinctorius spores 

at outplanting time proves ineffective. Canadian Journal of 

Forest Research 13: 1021-1023.

Alvarez, I.F.; Trappe, J.M. 1983b. Effects of application rate and 

cold soaking pretreatment of Pisolithus spores on effectiveness 

as nursery inoculum on western conifers. Canadian Journal of 

Forest Research 13: 533-537.

Barnes, L.R.; Johnson, C.R. 1986. Evaluation of ericoid 

mycorrhizae and media on establishment of micropropagated 

Rhododendron chapmanii Gray. Journal of Environmental 

Horticulture 4: 109-111.

Barnett, J.P. 1982. Relating field performance of containerized 

longleaf and shortleaf pine seedlings to mycorrhizal 

inoculation and initial size. In: Proceedings, 7th North 

American Forest Biology Workshop; 1982 July 26-28; 

Lexington, KY: 358-367.

Baser, C.M.; Garrett, H.E.; Mitchell, R.J.; Cox, G.S.; Starbuck, C.J. 

1987. Indolebutyric acid and ectomycorrhizal inoculation 

increase lateral root initiation and development of 

container-grown black oak seedlings. Canadian Journal of 

Forest Research 17: 36-39.

Beckjord, P.R.; McIntosh, M.S. 1984. Growth and fungal 

persistence by Quercus rubra inoculated with ectomycorrhizal 

fungi and planted on a clear-cutting and strip mine. Canadian 

Journal of Botany 63: 15 71-15 74.

Beckjord, P.R.; Melhuish, J.H., Jr.; Hacskaylo, E. 1986. 

Ectomycorrhiza formation on sawtooth oak by inoculation 

with basidiospore chips of Pisolithus tinctorius and 

Scleroderma citrinum. journal of Environmental Horticulture 

4: 127-129.

Beckjord, P.; Melhuish, J., Jr.; McIntosh, M. 1985. Influence of 

nitrogen and phosphorus fertilization on ectomycorrhizal 

formation of Quercus alba and Q. rubra seedlings by 

Pisolithus tinctorius and Scleroderma auranteum. In: Molina, 

R. (ed.). 6th North American Conference on Mycorrhizae. 

1984 June 25-29. Bend, OR. Corvallis, OR: Forest Research 

Laboratory: 221.

Benson, D.A.; lyer, J.G. 1978. Ectomycorrhizas and quality of 

nursery stock. Tree Planters' Notes 29: 3-7.

160
Dixon, R.K.; Garrett, H.E.; Cox, G.S.; Marx, D.H.; Sander, I.L. 

1984a. Inoculation of three Quercus species with eleven 

isolates of ectomycorrhizal fungi. I. Inoculation success and 

seedling growth relationships. Forest Science 30: 364-372.

Dixon, R.K.; Garrett, H.E.; Cox, G.S.; Pallardy, S.G. 1984b. 

Mycorrhizae and reforestation success in the oak-hickory 

region. In: Duryea, M.L.; Brown, G.N. (eds.). Seedling 

physiology and reforestation success. Dordrecht, The 

Netherlands: Martinus Nijhoff: 301-319.

Dixon, R.K.; Pallardy, S.G.; Garrett, H.E.; Cox, G.S.; Sander, 

I.L. 1983. Comparative water relations of container-grown 

and bare-root ectomycorrhizal and nonmycorrhizal Quercus 

velutina seedlings. Canadian Journal of Botany 61: 

1559-1565.

Donald, D.G.M. 1975. Mycorrhizal inoculation for pines. South 

African Forestry Journal 92: 27-29.

Dong, H.; Burdett, A.N. 1986. Chemical root pruning of Chinese 

pine seedlings raised in cupric sulfide impregnated paper 

containers. New Forests 1: 67-73.

Duddridge, J.A.; Malibari, A.S.; Read, D.J. 1980. Structure and 

function of mycorrhizal rhizomorphs with special reference to 

their role in water transport. Nature (London) 287: 834-836.

Ekwebelam, S.A.; Odeyinde, M.A. 1985. Field response of Pinus 

species inoculated with ectomycorrhizal fungi in Nigeria. In: 

Molina, R. (ed.). 6th North American Conference on 

Mycorrhizae; 1984 June 25-29; Bend, OR. Corvallis, OR: Forest 

Research Laboratory: 220.

Ekwebelam, S.A.; Reid, C.P.P. 1983. Effect of light, nitrogen 

fertilization, and mycorrhizal fungi on growth and 

photosynthesis of lodgepole pine seedlings. Canadian 

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Index of Common and

Scientific Names

Trees

alder

American green alder Alnus crispa (Ait.) Pursh 80

European alder A. glutinosa (L.) Gaertn. 141, 144

ash Fraxinus spp. 104

baldcypress Taxodium spp. 105

basswood Tilia spp. 105

bearberry Arctostaphylos uva-ursi (L.)

Sprengel 143, 144

beech Fagus spp. 105

birch

yellow birch Betula alleghaniensis Britton 118,

120, 141, 144

sweet birch B. lenta L. 141, 144

cedar

Atlas cedar Cedrus atlantica (Endl.) Manetti ex

Carr. 141, 144

"cedar"

Alaska-cedar Chamaecyparis nootkatensis (D.

Don) 52, 74, 105

incense-cedar Libocedrus decurrens Torr. 80, 105

northern white-cedar Thuia occidentalis L. 64

western redcedar T. plicata Donn. 124, 137

cherry Prunus spp. 104

Douglas-fir Pseudotsuga menziesii (Mirb.)

Franco 14, 23, 26, 27, 29, 38, 45, 48, 52, 56, 58,

60, 74, 75, 79, 80, 104, 105, 109, 1 1 1, 1 12, 1 15, 1 16,

119, 121, 122, 123, 125, 128-130, 133, 134, 136,

137, 139, 140-142, 146, 148, 149, 151, 153-155

eucalyptus

river redgum eucalyptus Eucalyptus camaldulensis

Dehnh. 140-143

fir

grand fir Abies grandis (Dougl. ex. D. Don)

Lindl. 23, 26, 29

noble fir A. procera Rehd. 52, 74

subalpine fir A. lasiocarpa (Hook.) Nutt. 26

white fir A. concolor (Gord. & Glend.) Lindl. ex

Hildebr. 80

giant sequoia Sequoiadendron giganteum

(Lindl.) 52, 74

hazel Corylus spp. 105, 107

hemlock

eastern hemlock Tsuga canadensis (L.) Carr. 142,

147

mountain hemlock T. mertensiana (Bong.) Carr 52,

74

western hemlock T. heterophylla (Raf.) Sarg. 23,

35, 52, 56, 63, 66, 74-75, 1 1 1, 1 19, 121, 126, 136,

140, 141, 147

holly Ilex sp. 138

juniper Juniperus spp. 104, 124

larch

European Larch Larix decidua Mill. 64, 148

Japanese larch L. kaempferi (Lambert) Carr. 64

Tamarack L. laricina (Du Roi) K. Koch. 64, 140,

141, 144

western larch L. occidentalis Nutt. 26, 29, 52, 55,

66, 74, 1 18, 141, 142, 144

magnolia

southern magnolia Magnolia grandiflora L. 153

maple Acer spp. 104, 105, 124

168
169

poplar, cottonwood Populus spp. 104, 105, 142,

146

redwood Sequoia sempervirens (D. Don) Endl. 52,

74, 124

spruce

black spruce Picea mariana (Mill.) B.S.P. 23, 64, 

118, 140, 144

blue spruce P. pungens Engelm. 30, 52, 74, 118, 

126

Engelmann spruce P. engelmannii Parry ex Engelm. 

23, 29, 32, 52, 74, 80, 112, 118, 121, 125, 134, 141, 144

Norway spruce P. abies (L.) Karst. 23, 141, 144

red spruce P. rubens Sarg. 118

Sitka spruce P. sitchensis (Bong.) Carr. 32, 40, 

140, 141, 143, 144, 148

white spruce P. glauca (Moench) Voss 23, 32, 

118, 121, 140, 144

Chapman rhododendron Rhodendendron chapmanii

Gray 141, 147

sweetgum Liquidambar spp. 104, 124

sycamore Platanus spp. 104, 124

walnut Juglans spp. 104

willow Salix spp. 105

yellow-poplar Liriodendron tulipifera L. 105, 153

Oak

black oak Quercus velutina Lam. 142, 143, 147 

bur oak Q. macrocarpa Michx. 136, 142, 147 

English oak Q. robur L. 140, 142, 143, 147 

laurel oak Q. laurifolia Michx. 119, 120 

northern red oak Q. rubra L. 140, 142, 143 

sawtooth oak Q. acutissima Carruth. 142, 143, 147 

white oak Q. alba L. 142, 143, 147

pecan Carya illinoensis (Wangenh.) K. Koch 141,

144, 155

pine

Aleppo pine Pinus halepensis Miller 142, 145

Austrian pine P. nigra Arnold 119, 120, 126, 142,

145

Caribbean pine P. caribaea Mill. 141-143, 145

Chinese pine P. tabuliformis Carr. 152

eastern white pine P. strobus L. 23, 64, 119, 142,

146, 152

jack pine P. banksiana Lamb. 23, 30, 31, 64, 119,

120, 140, 141, 143, 145

jeffrey pine P. jeffreyi Grev. & Balf. 78

limber pine P. flexilis James 22, 142, 145

loblolly pine P. taeda L. 58, 142, 146, 152, 156

lodgepole pine P. contorta Dougl. ex Loud. 23,

55, 56, 74, 80, 106, 1 1 1, 1 19, 123, 124, 140-142,

148, 152

longleaf pine P. palustris Mill. 26, 56, 80, 119,

120, 142, 145, 146, 152

maritime pine P. pinaster Aiton 140-143, 146

Monterey pine P. radiata D. Don 140-143, 146

Pinyon P. edulis Engelm. 21, 27

oocarpa pine P. oocarpa Schiede 142, 145

ponderosa pine P. ponderosa Dougl. ex Laws. 24,

26, 27, 29, 74, 80, 106, 112, 119-121, 125, 134,

140-143, 146, 148, 152, 156

red pine P. resinosa Ait. 23, 30, 31, 64

sand pine P. clausa (Chapm. ex Engelm.) Vasey ex

Sarg. 141, 145

Scotch pine P. sylvestris L. 23, 52, 56, 70, 74

shortleaf pine P. echinata Mill. 68, 80

slash pine P. elliottii Engelm. 80, 142, 145

sugar pine P. lambertiana Doug]. 21

Taiwan red pine P. taiwanensis Hayata 140, 141,

142, 146

Virginia pine P. virginiana Mill. 142, 146

western white pine P. monticola Dougl. ex D.

Don 15, 48, 89, 119, 120, 140-142, 145
Insects and Related Organisms

aphids

balsam wooly adelgid (aphid) Adelges picea

(Ratzeburg) 78, 89

giant conifer aphids Cinara spp. 56, 57, 89

root aphid Rhizomaria piceae (Hartig) 34, 48, 49

cutworms Euxoa spp. 6, 31-32, 89

variegated cutworm Peridroma saucia (Hubner) 89

European pine shoot moth Rhyacionia buoliana

(Denis & Schiffermuller) 77

flies

European crane fly (marsh crane fly) Tipula paludosa

Meigen 6, 34, 40, 41, 77, 90

dark-winged fungus gnats (Sciaridae) Bradysia

spp. 6, 34, 42, 43, 89

shore flies (Ephyridae) 42, 43

greenhouse whitefly Trialeurodes vaporariorum

Westwood 6, 44, 58

lygus bugs

Lygus hespurus (Knight) 58

 tarnished plant bug L. lineolaris (Palisot de

Beauvois) 58, 59

spider mites (Tetranychidae) 6, 44, 57, 64

thrips 60

weevils

black vine weevil Otiorhynchus sulcatus

Fabricius 34-37, 89

strawberry root weevil O. ovatus L. 34-37, 89

webworms Chrysoteuchia topiaria (Zeller) 34, 38,

39

Crambus spp. 38, 39

Pests

Disease Fungi

Botrytis cinerea Pers.:Fr. 9, 51-55, 61, 74-76, 81,

85-89

Caloscypha fulgens (Pers.) Boudier 22, 23, 78

Collectotrichum acutatum Simmonds 56

Cronartium fusiforme Hedgcock & Hunt ex

Cummins 56

Cylindrocarpon spp. 48

Fusarium spp. 24, 26, 28, 44-46, 48, 50, 51, 54,

78-81, 85, 87-89

F. avenaceum (Fr.) Sacc.26, 44

F. moniliforme Sheldon 26

F. oxysporum Schlecht. 26, 27, 44, 85, 116

F. roseum Lk.:Fr. 26

F. solani (Mart.) Appel & Wollenw. 27, 44

F. tricinctum (Corda) Sacc. 27

Phytophthora spp. 26, 44, 46, 47, 81, 84, 86, 88,

89

P. cinamoni Rands 116

Pythium spp. 26, 27, 46, 47, 81, 84, 86-89

Rhizoctonia spp 27, 44, 56, 81, 86, 88, 89

Sirococcus. spp. 17, 19, 87

S. strobilinus Preuss. 17, 32, 33

Sphaeropsis spp. 17, 19

S. sapinea (Fr.) Dyko & Sutton 17

Tricoderma spp. 85

Bacteria

Bacillus spp. 85

Pseudomonas spp.85

Plants

algae 66-68, 82, 88

bitterbrush 30

bittercress Cardamine pennsylvanica 64

bryophytes 67

lichens 67

liverworts 64, 66-68, 82, 88

moss 66-68, 82, 88

Oregon-grape Berberis aquifolium Pursh. 138

sorrel Oxalis spp. 64

Animals

goldfinches Carduellis spp. 22

meadow vole Microtus pennsylvanicum 73

pine vole M. pinetorum 73

170
Mycorrhizal fungi

Acaulospora spp. 105, 114

Alpova trappei Fogel 111

Amanita muscaria (L.:Fr.) Hooker 106, 140, 144

Amphinema byssoides (Pers.:Fr.) J. Erikss. 121, 125

Astraeus hygrometricus (Pers.) Morgan 140, 145

Balsamia spp. 105

Boletus satanus Lenz 106

Cenococcum geophilum Fr. 104, 105, 119, 121,

125, 126, 129, 139, 140, 144-147

Chamonixia caespitosa Roll. 131

Chloridium spp. 105

Cortinarius spp. 105

Elaphomyces spp. 105

Endogone lactiflua Berk. & Br. 120, 140, 146

Entrophospora spp. 105, 114

Gastroboletus turbinatus (Snell) Smith & Singer 108

Gautieria spp. 105

Geopora spp. 105

Genea spp. 105

Gigaspora spp. 105, 114

Glomus fasciculatum (Thaxter) Gerdemann &

Trappe 114

G. microcarpum Tul. & Tul. 114

Hebeloma arenosa Burdsall, McFall & Albers 125,

126

H. crustuliniforme (Bull.) Quel. 106, 111, 125, 129,

135, 136, 139, 140, 144, 146, 148, 156

H. cylindrosporum Romagn. 140, 144

H. sinapizans (Paulet:Fr.) Gillet 140, 146

Hydnangium carneum Wallroth 140, 144

Hydnotra spp. 105

Hymenogaster sp. 106,- 132

Hysterangium spp. 105

Inocybe lacera (Fr.) Kummer 125

Laccaria bicolor (Maire) Orton 140, 144

L. laccata (Scop.:Fr.) M.C. Cooke 112, 116, 120,

121, 125, 129, 134-136, 139, 141

L. Proxima (Boud.) Pat. 141, 145

Lactarius paradoxus Beardslee & Burl. 141, 145

L. rubrilacteus Hesler & Smith 123

Leucopaxillus ceralis (Lasch) Singer 116

Leccinum spp. 105

Martellia medlockii Trappe & Castellano 106, 123

Mycelium radicis atrovirens = Phialocephala

dimorphospora Kendrick 125

Paxillus involutus (Batsch:Fr.) Fr. 141, 146

Pezizella ericae Read 141, 147

Phialophora spp. 105

Pisolithus tinctorius (Pers.) Coker & Couch 120,

128, 129, 134-136, 139, 141, 142, 145-149, 152,

153, 156, 157

Rhizopogon colossus A.H. Smith 142, 146

R. evadens A.H. Smith 134

R. fuscorubens A.H. Smith 134

R. luteolus Fries & Nordholm 129, 134, 142, 145,

146

R. nigrescens Coker & Couch 142, 145

R. occidentalis Zeller & Dodge 108

R. ochraceorubens A.H. Smith 134

R. ochraceisporus A.H. Smith 131

R. parksii A.H. Smith 134

R. roseolus (Corda) Th. M. Fires 129, 142, 146

R. rubescens Jul. & Tul.) Tul. & Tul. 125, 126,

142, 146

R. smithii Hosford 108

R. subgelatinosus A.H. Smith 134

R. truncatus Linder 131, 134

R. villosulus Zeller 134

R. vinicolor A.H. Smith 1 12, 1 15, 128, 129,

133-134

Russula spp. 105

Sclerocystis sp. 105, 114

Scleroderma aurantium Pers. 143, 147

S. bovista Fr. 143, 145

S. cepa Pers. 106

S. citrinum Pers. 143, 147

S. verrucosum Pers. 143, 144

S. paradoxum Beaton 143, 144

S. texense Berk. 143, 145

Scutellospora spp. 105, 114

Sphaerosporella brunnea (Albertini & Schwein.:Fr.)

Surcek & Kubicka 121, 143, 145

Suillus bovinus (L.:Fr.) O. Kuntze 129

S. granulatus (L.:Fr.) O. Kuntze 143, 145-148, 152

S. luteus (L.:Fr.) S.F. Gray 129, 143, 147

S. tomentosus (Kauffm.) Singer, Snell & Dick 143,

145

Thelephora terrestris Ehrhart:Fr. 104, 109,

120-122, 129, 139, 143-147, 156

Tricholoma spp. 104

Tuber gibbosum Harkness 131, 132

171