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1978 Nitrogen fixation, photosynthesis and early growth of Alnus glutinosa Jeffrey Owen Dawson Iowa State University

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DAWSON, JEFFREY 0.s'£M WITROGE'n FIXATION, PhUTOSY"T*E?iS EA«LY GROWTH OF ALNuS GLUTlnCSA.

IOWA STATt. UNIVERSITY, PH.D., 1978

Universitv Micrdfilnris International 300ZTTE «OAD. ABBOH, MI-JSIOS Nitrogen fixation, photosynthesis

and early growth of Alnus glutinosa

by

Jeffrey Owen Dawson

A Dissertation Submitted to the

Graduate Faculty in Partial Fulfillment of

The Requirements for the Degree of

DOCTOR OF PHILOSOPHY

Department: Forestry Major: Forestry (Biology)

Signature was redacted for privacy.

!ri Charge of M^jor work

Signature was redacted for privacy.

For the Major Department

Signature was redacted for privacy.

F( uate College

Iowa State University Ames, Iowa

1978 ii

TABLE OF CONTENTS

INTRODUCTION

LITERATURE REVIEW

The Host Plant

The Endophyte

OBJECTIVES

PROCEDURES

Plant Material

Nutrient Culture System

Acetylene-Reduction Assay

Photosynthesis

Tissue Analysis

RESULTS

DISCUSSION

ACKNOWLEDGMENTS

LITERATURE CITED

APPENDIX iii

LIST OF TABLES

Page

Table 1. Growth of selected Alnus glutinosa cuttings under uniform conditions 29

Table 2. Nitrogen-free Van der Crone's solution (half-strength) 32

Table 3. Analysis of variance from regression on dependent variable leaf area 43

Table 4. Analysis of variance from regression on dependent variable nitrogenase activity 43

Table 5. Total plant nitrogen (mg) at harvest 46

Table 6. Percentage nitrogen in alder tissues (mean values and 95 percent confidence intervals) at harvest 49

Table 7. Shoot/ratios at harvest 50

Table 8. Mean squares from analysis of variance for results of experiments with fertilized alder clones 51

Table 9. Mean squares from analysis of variance for results of experiments with nodulated alder clones 52

Table 10. Mean squares from analysis of variance for results of experiments with uninoculated alder clones 52

Table 11. Linear correlation coefficients and probability levels for relationships between mean clonal values of variables determined at time of harvest 55

Table A--1. Data on inoculated alders in hydroponics 77

Table A--2. Data on inoculated alders in hydroponics 78

Table A-•3. Data on inoculated alders in hydroponics 79

Table A-•4. Data on inoculated alders in hydroponics 80

Table A--5. Data on inoculated alders in hydroponics 81

Table A-•6. Data on inoculated alders in hydroponics 82 iv

Page

Table A-7. Data on inoculated alders in hydroponics 83

Table A-8. Data on inoculated alders in hydroponics 84

Table A-9. Data on inoculated alders in hydroponics 85

Table A-10. Data on uninoculated alders in hydroponics 86

Table A-11. Data on uninoculated alders in hydroponics 87

Table A-12. Data on u^.noculated alders in hydroponics 88

Table A-13. Data on uninoculated alders in hydroponics 89

Table A-14. Data on uninoculated alders in hydroponics 90

Table A-15. Data on uninoculated alders in hydroponics 91

Table A-16. Data on uninoculated alders in hydroponics 92

Table A-17. Data on fertilized alders in potting mix 93

Table A-18. Data on fertilized alders in potting mix 94

Table A-19. Data on fertilized alders in potting mix 95

Table A-20. Data for regression equations to estimate initial dry weight and nitrogen content of alders 96

Table A-21. Data for regression equations to estimate alder leaf area 97 V

LIST OF FIGURES

Page

Figure 1. Diurnal fluctuation in acetylene reduction by whole, nodulated root systems of Alnus glutinosa 39

Figure 2. Regression of late-afternoon acetylene reduction rate on leaf area of nodulated Alnus glutinosa 47

Figure 3. Regression of photosynthesis rate on total- plant nitrogen of nodulated Alnus glutinosa at time of harvest 48

Figure 4. Clonal means for dry mass and percentage nitrogen of fertilized Alnus glutinosa plants at time of harvest 53 1

INTRODUCTION

Most plants and animals in the world suffer from nitrogen

deficiency at some time during their existence. Terrestrial

plants and animals suffer nitrogen deficiency while they are bathed

in an atmosphere that is approximately 80 percent nitrogen.

The ability to convert gaseous nitrogen into a form which

plants and animals can use is restricted to a few bacteria,

algae and actinomycetes. The most efficient of these prokaryotic

nitrogen fixers have a ready supply of metabolites for carrying on

nitrogen fixation. For instance, blue-green algae have an endogenous

supply of.photosynthate, while some bacteria and actinomycetes in root

nodules are supplied with photosynthetic products by a host plant.

Delwiche (1970) points out the increasingly important role that

industrial manufacture of nitrogen fertilizer, using the Haber process,

plays in the world's nitrogen cycle. Industrially fixed nitrogen is

used extensively to increase yields of food and fiber from cultivated

plants.

The energy costs of industrial nitrogen fixation are considerable.

The manufacture and application of 100 pounds of nitrogen as urea

are estimated to have a total energy cost of 50 gallons of #4 fuel

oil (Smith and Johnson, 1977), making nitrogen fertilization, aside from harvesting, the largest potential consumer of energy in silviculture. Second in potential energy consumption is mechanical

site preparation for planting. 2

Biological nitrogen fixation is a process that derives its

energy directly from the sun. The energy costs of biological

nitrogen fixation are also considerable. To convert one molecule

of dinitrogen into two ammonia molecules requires 15 ATP (Conn and

Stumpf, 1972). However, from a consideration of the thermodynamics

of nitrogen fixation, Bergersen (1971b) concluded that the energy

requirements for biological nitrogen fixation and the assimilation

of combined nitrogen may be similar. If this is true, then a

terrestrial plant might expend about the same amount of energy in fixing atmospheric nitrogen symbiotically as it expends taking up

combined nitrogen from the soil in the usual manner, making the energy cost of biological nitrogen fixation negligible.

One general goal of this study was to enhance N-fixation in the symbiotic system of European alder, Alnus glutinosa (L.) Gaertn., and an actinomycete-like endophyte. The simplest way to improve nitrogen-fixing efficiency is by genetic screening of both hosts and endophytes to determine the combinations best suited for a particular environment (Brill, 1977). But in the case of alder, genetic screening is limited to the host plant because the nitrogen- fixing endophyte of alder has not yet been reproducibly isolated in virulent form. However, vegetative propagation of alder by stem cuttings makes clonal, host-plant material readily available for selection work. So, in order to assess the effects of host-genotype on early growth, the response of clonal European alder to nitrogen J

supplied by a North American endophyte, to combined nitrogen and

to nitrogen starvation under controlled conditions was studied.

Indicators of photosynthesis and nitrogen fixation of clonal Euoropean

alder were also studied to determine relationships between photo­

synthesis and nitrogen fixation for specific combinations of host

plants, inoculum and environment.

The information obtained should be useful in establishing selection procedures for increasing symbiotic nitrogen fixation in plant systems and in identifying clonal lines of alder with the potential for increased nitrogen fixation. 4

LITERATURE REVIEW

The Host Plant

Named for the stickiness of its buds and twigs, A. glutinosa or

European alder is a native of Europe, northern Africa and Asia. It

is also naturalized locally in parts of eastern Canada and the north­

eastern United States.

This medium sized tree has a form ranging from strongly excurrent

to distinctly shrubby (Funk, 1965). It has been reported to reach a

maximum height of 35 m (Fernald, 1950), a maximum diameter of about

1 m and a maximum age of about 120 years (McVean, 1953a) on the best

sites in Europe. In Iowa, a European alder near the administration

building of the Mental Health Institute at Independence was 20 m tall and .76 m in diameter at breast height when measured in July, 1976.

In Britain A- glutinosa is a species of stream and lake sides as well as of poorly drained soils (McVean, 1953a). It also does well on intermittently moist sites if rainfall is adequate (Loycke, 1963).

McVean (1956a) states that a minimum of 50 to 65 cm of rainfall per year are necessary for seedling establishment, except where the surface soil falls within the capillary fringe of the water table.

Natural reproduction has occurred in the Des Moines River valley at the 4-H camp near Madrid, Iowa. Seedlings were observed in 1977 near a plantation which receives an average of 80 cm of rainfall per year. Naturalized stands of A. glutinosa also occur in the DuPage

County forest preserve in northern Illinois. 5

In Eurasia the southeastern boundary of European alder distribution

corresponds closely with the 20-inch (50.8 cm) annual rainfall line

(McVean, 195 3a). It does not extend north into regions that have less

than six months in which the mean daily temperature is above freezing

and is susceptible to late spring frost (Ulrich, 1962). However,

Groszman and Melzer (1933), as cited in McVean (1953a), reported that

the species is hardy to winter temperatures of -65°F.

Being a pioneer and opportunistic species A. glutinosa is

capable of colonizing even the rawest soil material, including road

banks in the west of Scotland (McVean, 1956c). It is more shade-

tolerant than willow, poplar, birch, or Scotch pine, but less shade-

tolerant than eastern white pine or Douglas fir (Leibundgut, 1963).

Natural European alder communities often include ash, birch, willow,

and oak as associates (McVean, 1953a).

European alder is monoecious and some trees flower as early as

the beginning of their fourth growing season (Funk, 1965). Flowers

of both sexes develop from buds that begin to develop in the spring

of each year and pollination occurs in the early spring of the next year. After pollination, fertilization does not take place until late July or early August (McVean, 1955a). Seed begins to fall in late September of early October (Ulrich, 1962).

Seed crops are generally heavy (Loycke, 1963) and the viability of cross-pollinated seed may range up to 90 percent (McVean, 1955a). 6

Seeds have no dormancy requirements and few remain viable beyond the

first germination season (McVean, 1955b). Optimal seed germination

requires a temperature of 26°C, and the elongating radicle is par­

ticularly sensitive to drying and cold (McVean, 1955a). This provides

a partial explanation of European alder's hydrophytic behavior.

Seeds possess air bladders enabling them to float. Because they are small and wingless, they usually are not spread far where wind is the only means of dispersal. McVean (1955b) concluded that water is the principal means of seed dispersal.

European alder is easily coppiced and reproduces rapidly from stump sprouts, though root suckers are rare (McVean, 1953a). The root system is extensive, possessing both surface and deep branches with considerable plasticity of structure, enabling the tree to sur­ vive on either water-logged soils or those of deep water table

(McVean, 1953a). The shallow, fibrous roots are nodulated and infected by mycorrhizal fungi. Development of nodules and mycorrhizae is thought to have little effect on establishment except on the most infertile sands (McVean, 1955a).

Optimal growth occurs at soil pH of approximately 5.4 (Bond et al., 1954). Alkaline conditions reduce growth but are better for nodulation which is optimal at soil pH around 7.0.

Because of its broad range, A. glutinosa might be expected to exhibit much genotypic variation. Kjersgaard (1963), as cited in

Funk (1965), reported results of a study which lasted 20 years and 7

included several Danish provenances. He concluded that several local

races could be distinguished with slightly varying disease resistance,

but growth differences were inconsequential. McVean (1953b) found

differences in the date of growth initiation in the spring and time

of leaf fall in autumn between seedlings of Scottish origin and seed­

lings from further south in England and Wales. He also observed clinal

variation in leaf size and pistillate catkin dimensions of British

alders. Funk (1965) reported that four southern Bavarian provenances

were visibly taller as a group than provenances from central Germany,

northern Germany, and central Sweden both after one year in a nursery

and two subsequent years on strip-mined land in Ohio. A range-wide

provenance study of A. glutinosa for the north-central United States

is now being conducted by Richard B. Hall of the Forestry Department

at Iowa State University.

A. glutinosa hybridizes readily with many other alders.

Particularly vigorous hybrids have been reported for A. cordata x

A. glutinosa (Krussman, 1956), A. glutinosa x A. incana (Kobendza,

1956) and A. glutinosa x A. rubra (Ljunger, 1959).

Spalding (1958) lists three diseases of European alder in the

United States, all of minor importance: Phymatotrichum omnivorum, a root rot of alder growing on specific calcareous soils in Texas;

Polyporous versicolor, a saprophytic sap-rot that can become parasitic; and Septoria-alni, a leaf spot reported from Wisconsin. 8

The poplar and willow borer, Cryptorhynchus laputhi, can cause

serious damage by girdling and boring at the bark of alders in the

midwestern United States (Funk, 1965). A host of other fungi and

insect pests, including the cottonwood leaf beetle, are associated

with European alder (Funk, 1965).

Succulent shoots are browsed by cattle and deer but not

preferentially (McVean, 1953a). Rabbits have caused some damage to young plantations in Ohio (Funk, 1965).

A. glutinosa is the only tree suitable for wood production in marshy areas of Europe where dominant stands can produce straight stems without branching due to natural pruning (Scamoni, 1960). Under marshy conditions 60 to 90 year old stands can yield 507 to 718 of stemwood per ha (Wiedemann and Schober, 1957). The wood is suitable for furniture, plywood, clogs, hat blocks, barrel staves and small turnery work. A very high grade of charcoal is produced from alder wood. All parts of the plant, particularly the bark, are rich in tannins.

European alder is also useful to man because of its ability to

symbiotically fix atmospheric nitrogen. Dinger (1895) was among

the first to publish experimental results relating A. glutinosa to

nitrogen fixation. His experiments with alder on a bare, sandy soil

in the Netherlands showed that plants with the most nodules grew best

and had higher leaf nitrogen contents than other plants on the same

soil. He demonstrated that removing nodules slowed leaf expansion and lowered nitrogen content. Hiltner (1896) and Nobbe and Hiltner 9

(1904) concluded that a fixation of elementary atmospheric nitrogen

occurred in A. glutinosa nodules owing to the fact that nodule-

bearing alder can be grown in nitrogen-free nutrient solution.

Krebber (1932) and Roberg (1934) in repeating these experiments

convincingly demonstrated excellent growth of plants with nodules

on nitrogen-free nutrient solutions. Kjeldahl analysis of fixed

nitrogen was demonstrated by von Plotho (1941). Burris and Miller

(1941) established that assimilation of was definitive proof

for nitrogen fixation. Discovery of the acetylene reduction assay

for nitrogenase activity provided increased sensitivity over the

method (Dilworth, 1966 and Schollhorn and Burris, 1966). Both

methods have subsequently been successfully applied to A. plutinosa

(Bond, 1955 and Akkermans, 1971).

Because of their nitrogen fixing symbioses, European alder and others among the 157 species in 13 genera of non-leguminous angiosperms with Alnus-type nodules (Bond, 1976) contribute significantly to the nitrogen economy of a number of ecosystems (Bond, 1974).

Living European alder nodules excrete albuminoid compounds which might be a mechanism of nitrogen enrichment (Virtanen and

Saastamoinen. 1936). The decay of old nodules and roots might also add nitrogen to the soil (Burris, 1959). Wollum and Youngberg (1964) demonstrated that both litter and the subterranean parts of clipped

Alnus rubra and Ceanothus velutinus had a beneficial effect on the growth of Pinus radiata. The increase in soil nitrogen under European 10

alder may, however, result more from decomposition of enriched leaf

litter than from direct excretion by root nodules (Bond, 1955; Bond,

1956a; Mikola, 1958; Redko, 1958; and Virtanen, 1957).

Nitrogen accumulations ranging from 60 to 209 kg per ha per year

have been estimated for Alnus spp. (Tarrant, 1968). Annual nitrogen

return was from 80 to 200 kg per ha per year during the last seven

years of growth in a 30-year-old A. rubra stand (Gessel and Turner,

1974). About 85 kg per ha per year of nitrogen were added to an

Alnus rugosa ecosystem in which the alder plants had an average age

of 16 years (Voight and Steucek, 1969). About 93 percent of the total nitrogen in the ecosystem was in the soil. More nitrogen was accumu­

lated on sites with moister soils. Of the nitrogen in the alder

plants, about 75 percent was found in the stems and twigs, but the highest concentrations of nitrogen were found in leaves and nodules.

Thickets of Alnus crispa five years old and 1.5 m tall were calculated to add 157 kg per ha per year nitrogen to the ecosystem by leaf fall alone (Lawrence, 1958). A. crispa plays a major role in soil forma­ tion on recently deglaciated sites in North America by adding nitrogen and organic matter as well as by acidifying the upper layers of glacial till (Crocker and Major, 1955). Populus trichocarpa saplings associated with A. crispa weighed 22.5 times as much as those of equal age in non- alder areas (Lawrence, 1953).

The economic significance of A. glutinosa in European forestry for interplanting Pinus sylvestris, Picea abies, and Populus spp. has long been recognized (Stone, 1955; Virtanen, 1957; Redko, 1958; and 11

Van der Meiden, 1950). Nikola (1958) discovered that leaf litter of

Alnus incana and A. glutinosa were superior to those of other trees

for the growth of Pinus sylvestris. The distance of apple trees

from A. glutinosa hedges and leaf nitrogen content of the apple leaves

were negatively correlated in unfertilized plots in the Netherlands

(Delver and Post, 1968). The U.S. Forest Service is experimenting

with mixed stands of glutinosa and Juglans nigra. Mixed plantations

exist near Carbondale, Illinois, in the Amana Colonies of Iowa, and

at the 4-H camp in the Des Moines River valley near Madrid, Iowa.

Members of the following genera have also exhibited increased growth

when grown in association with Alnus spp.: Fraxinus, Liquidambar,

Platanus.. and Pseudotsuga (Tarrant, 1968).

Because of European alder's ability to improve soil, it has been

employed widely in rehabilitating mine spoils in Great Britain, the

Netherlands, Germany, and the United States. It is routinely used

to stabilize flood deposits and landslide areas in Rumania,

Czechoslovakia and Germany (Tarrant, 1968). Other areas in the

northern hemisphere where Alnus species have been used to improve

soil fertility are the U.S.S.R., Italy, Denmark, Japan, and Taiwan

(Kohnke, 1941; Tarrant, 1968). European alder is also planted widely

as an ornamental in the eastern United States.

The Endophyte

Nitrogenase, the enzyme responsible for nitrogen fixation, is synthesized by the prokaryotic, microbial symbiont in the nodules of 12

A. glutinosa (Skeffington and Stewart, 1976). Characters of this

prokaryotic, microbial endophyte suggest that it is an actinomycete

(Becking et al., 1954; Gardner, 1965; and Lalonde and Knowles, 1975b).

Nodulated European alder growing in nitrogen-free media has special micronutrient requirements. Cobalt, molybdenum and copper are necessary and are implicated in nitrogenase-mediated nitrogen fixation. Cobalt is required as a micronutrient in several non- leguminous root nodule plants (Hewitt and Bond, 1966). Severe nitrogen deficiency develops in A. glutinosa plants grown without cobalt (Bond and Hewitt, 1962). The necessity for cobalt exists in the nodules, and its deficiency causes nodules to fail to fix nitrogen at the normal rate (Hewitt and Bond, 1966). Cobalt is contained in vitamin

The actinomycetes are active producers of vitamin analogues

(Bond et al., 1965). Vitamin B^g analogues are much more abundant in nodules as compared with roots (Bond e^ aJ^., 1965), supporting the belief that the endophyte is an actinomycete. Copper is an essential micronutrient for higher plants as well as for many microorganisms including actinomycetes (Waksman, 1959). It is required for nitrogen fixation in nodulated A. glutinosa (Bond and Hewitt, 1967). Molybdenum is an essential micronutrient for nitrogen fixation in A. glutinosa

(Hewitt and Bond, 1961). Molybdenum is concentrated in the nodular tissue of both molybdenum-deficient and molybdenum-supplied A. glutinosa

(Becking, 1962).

Nodules of A. rubra (Li et al., 1972) and A. glutinosa (Pizelle and Thiery, 1974) exhibit nitrate-reducing capability more than twice 13

that of root segments. Actinomycetes characteristically contain

nitrate reductase (Breed et al., 1957) and higher activities asso­

ciated with nodules are generally attributed to the actinomycetal

character of the endophyte.

Nodules of A. glutinosa also exhibit higher levels of plant

growth regulators than other parts of the plant. Indole acetic acid

(lAA) is the main auxin in root nodules of A. glutinosa, and, on a

fresh weight basis, is present in substantially larger amounts than

in roots (Dullhaart, 1970), perhaps representing a reaction by host

tissue to microbial infection, as in crown gall. Levels of gibberellin-

like activity are also higher in root nodules than in other parts of

young A. glutinosa plants (Henson and Wheeler, 1977).

Isolation in pure culture, identification and reinfection of

the host plant with causal organisms of root nodules in non-leguminous

angiosperms, including alders, has not been achieved (Becking, 1970a;

Quispel, 1974; Rogers and Wollum II, 1974; and Lalonde e£ , 1975).

Growth of the endophyte in culture needs to be such that cultures of

the endophyte can be produced in reasonable time and used to reinfect

alder roots to form nodules (Quispel, 1955). However, John Torrey of

the Cabot Foundation, Harvard University, Petersham, Massachusetts, has reported successful isolation in culture and reinfection of the host plant with the endophyte of Comptonia peregrina (personal communication,

August 19, 1977).

The loss of virulence of the alder endophyte in culture and the absence of European alder endophyte in some natural soils has lead 14

Becking (1970b) to suggest that the endophyte is an obligate or near-

obligate symbiont. Presently, an inoculum of crushed nodules is used

to infect European alder roots. However, soil samples taken from

sites in Britain where A. glutinosa had not grown for years produced

nodules on young alder plants, suggesting that the infectious bodies

of the endophytes can survive in soils for long periods, though not

grow, or that the endophyte can grow saprophytically in soils and,

thus, is not a near-obligate symbiont (Rodriguez-Barrueco, 1968).

In addition, Rossi (1964) demonstrated survival and multiplication

of the A. glutinosa endophyte in soil.

The endophyte that induces nodule formation in Ceanothus

velutinus in the western United States was still present in soil

that had not supported that plant for some 150 years (Wollum e£

al., 1968). But, the habitat soil of timber stands 200-350 years

old exhibited almost complete absence of infective Ceanothus endo­

phyte suggesting some decline.

Benecke (1969) found an endophyte capable of forming effective

nodules on Alnus viridis to be present in remote grassland topsoils

of New Zealand which contained only indigenous plant species. Native

beech-forest (Nothofagus) topsoil, on the other hand, had a microbial

population that did not regularly include nodule-forming organisms.

New Zealand's flora contains no indigenous alders and only two genera of non-legumes that form root nodules. The native Coriaria sarmentosa has nodules that were ineffective for the nodulation of A. viridis. 15

The decline or absence of non-leguminous endophytes in forest soil

while an endophyte present in a native New Zealand grassland topsoil

formed effective nodules on an exotic Alnus species suggests to me a

soil pH requirement for survival in soil of the actinomycete-like

endophyte in absence of specific hosts. Forest soil tends to be more

acidic than grassland soil and the soil pH optimal for nodulation

of at least one non-leguminous angiosperm is higher than optimal pH

for host plant growth (Bond e^ al., 1954). The endophytes which infect

alder must have infective forms in soil which are extraordinarily

ubiquitous in distribution.

European alder seeds sown in beds at the State Forest Nursery

in Ames, Iowa produce plants with nodulated roots even though no

native alders have occurred in the area in historic times. The only

native alder in Iowa is A. rugosa, which was common following the

retreat of Wisconsin glacial ice from Iowa 10,000 to 12,000 years ago.

It is now restricted to wet, cool sites near springs in northeastern

Iowa. Since there is no evidence that the alder endophyte is seed-

bom (Ferguson and Bond, 1953; Akkermans and Van Dijk, 1976), it is

likely that the endophyte used in this work originated in the mid- western United States. The importation and propagation of A. glutinosa

is effected primarily by seed.

The anatomy of alder nodules was described originally by Shibata

(1902) and later by Schaede (1933) using light microscopy. Nodules

are modified, dichotomously branched roots of arrested growth with a

coralloid appearance. In the nodules the endophyte occurs in three 16

main forms that have been ascribed different functions. The filamentous

form is the primary form to infect host cells; the vesicular form

may be the site of nitrogen fixation, while the bacteria-like form is

thought to be a resting stage concerned with reproduction and dispersal

(Becking, 1970b; and Akkermans, 1971).

Cross-inoculations using nodular material within the genus Alnus

show grades of incompatibility. Inter-generic crossing using alder

endophyte with other nodule-forming, non-leguminous angiosperms has

failed to produce nodules (Becking, 1970b). Geographic location of

members of the genus Alnus may have resulted in the evolution of

taxonomic affinities and consequent barriers to cross inoculations.

Mackintosh and Bond (1970) attempted 17 combinations of

nodular endophyte and host species within the genus Alnus. Out of

the 17 attempted, nine resulted in symbiosis. A. rugosa, for instance, was successfully inoculated with A. glutinosa nodules. Cross inocula­ tion of A. viridis with nodules of A. nepalensis and A. sinuata failed to produce nodules in New Zealand trials (Benecke, 1969). Immuno- labelling has been used to identify the A. crispa endophyte in nodules of successfully cross-inoculated A. glutinosa (Lalonde and Quispel,

1977). The endophyte apparently exists in a number of forms, each able to combine with a restricted number of exotic host species.

Schaede (1933) distinguished two types of nodules in A. glutinosa on the basis of the presence or absence of granula and suggested that they represent different strains of the endophyte. Van Dijk and Merkus

(1976) examined granula-rich and granula-free root nodules of 17

A. glutinosa by light and electron microscopy. Granula bore a strong

resemblance to spores of free-living actinomycetes. Healthy nodules

with granula (Gr ) possess 100 to 1000 times greater infective power

than healthy nodules without granula (Gr ), supporting the idea that

the granula or spores are infective particles (Akkermans and Van Dijk,

1976). Necrotic Gr nodules are 10 to 100 times more infective than

healthy Gr nodules, while necrotic Gr nodules are less infective than

healthy Gr^ nodules.

The nodular supply used in this study was assayed and found to

be exclusively Gr by Maurice Lalonde of the Kettering Laboratory,

Yellow Springs, Ohio (letter dated April 14, 1977). This supply of

nodules originated from nodules of A. glutinosa grown from seed at

the Iowa State Forest Nursery in Ames- They were used to inoculate

stock plants in nutrient solution and were propagated by repeated

harvest of nodules for infection of additional stock plants.

During infection, the free-living form of the actinomycete-like

endophyte of Alnus enters the plant through a root hair (Pommer, 1956

and Angulo-Carmona, 1974). There the hyphal endophyte perforates the

host cell wall and enters a cortical cell (Lalonde and DeVoe, 1975)

where it is surrounded by a membrane envelope of host origin (Lalonde and DeVoe, 1976). The host cell synthesizes, transports and deposits

pectic substances for the encapsulation of the intruder (Lalonde and

Knowles, 1975a). This pectic capsule has been observed in A. glutinosa

nodules (Lalonde and Knowles, 1975a). Bound enzymes seem to be randomly

distributed in the capsular pectic layers where they might facilitate

uptake or release of metabolites or function in the synthesis or the 18

degradation of the capsular pectic layers themselves (Lalonde and and DeVoe, 1976).

If the endophyte can degrade cell wall components with enzymes, then it must also be equipped to degrade its rigid pectate capsule

(Lalonde and Knowles, 1975a). Degradation of this pectic capsule could be a means by which the heterotryphic endophyte obtains carbo­ hydrate (e.g. galacturonic acid) from its autotrophic host (Smith et al., 1969). This carbohydrate could, then, be used by the endo­ phyte for nitrogen fixation.

In A. glutinosa, the vesicles of the endophyte are considered to be the main site of nitrogen fixation, and the tops of nodule lobes are the.primary location of nitrogen fixation activity within the nodule (Akkermans, 1971).

Nodules induced on Myrica fay a with the M. gale endophyte are small and fix no significant amount of nitrogen (Mian et al., 1976).

The finding that these nodules, lacking vesicles, are ineffective is consistent with the belief that vesicles are the main site of nitrogen fixation in Alnus-type nodules.

Most of the nitrogen fixed by A. glutinosa is converted to the amino acid citrulline (Miettinen and Virtanen, 1952) by a process that may involve the direct fixation of carbon dioxide in the root of the plant (Leaf, Gardner and Bond, 1958). Citrulline is the main vehicle for the translocation of nitrogen in the alder from roots to leaves via the transpiration stream (Gardner and Leaf, 1960; Bond, 1956b). 19

Both nitrogen fixation and nodulation are affected by a variety

of chemical and physical conditions including oxygen concentration,

substrate pH, combined nitrogen species, combined nitrogen concentration,

water availability, temperature, and light.

The nitrogen-fixing nodule requires 0^ for ATP production;

however, the 0^ sensitivity of nitrogenase dictates a requirement for

low pO^ inside the nodule (Bergersen, 1971a). A. glutinosa nodules have a higher 0^ requirement than roots (Ferguson, 1953); however, high 0^ concentration inhibits the nitrogenase-catalyzed reaction

(Becking, 1970b). Bond (1961) demonstrated that optimal nitrogen fixation in A. glutinosa occurred at a 12 percent oxygen concentration and that .40 percent oxygen virtually extinguished nitrogen fixation in root nodules. By bubbling gas mixtures of oxygen and nitrogen through vessels of nutrient solution, MacConnell (1959) found that inoculated root systems of A. glutinosa developed progressively fewer nodules as the oxygen level was lowered from the normal 21 percent. Nodulated plants were more sensitive to oxygen reduction than non-nodulated plants. Nitrogen fixation in legumes is also significantly improved by bubble aeration or by lowering the level of culture solution below the main zone of nodulation (Minchin and Pate, 1975). However, basic differences exist between Alnus-type nodules and legume nodules.

The endophyte in Alnus nodules is in cortical tissue outside a cylinder of vascular tissue. In legumes the bacterial endophyte is within a cylinder of vascular tissue and possesses leghaemoglobin to 20

aid in oxygen respiration. No haemoglobin has been found in Alnus

nodules (Virtanen, 1948), though a bound form of haemoglobin that is

not extractable from cell debris by aqueous solvents may exist in

alder (Davenport, 1960). Virtanen (1948) suggested that haemoglobin

may not be necessary in alder because of the endophyte's proximity

to the nodule surface where lenticels provide an opening through

superficial cork layers. These lenticels can be induced on stems and

roots of A. glutinosa by lowering oxygen tension and increasing carbon

dioxide concentration (McVean, 1956b). Their function may be to increase the efficiency of the plant's aeration system and assist respiration of the nodular endophyte. A possible internal aeration system might account for the reddish, oxidized deposit sheathing alder roots of all ages in reducing soils (McVean, 1956b). Such an aeration system would make A. glutinosa somewhat independent of oxygen concentration around the submerged parts of roots. Nodules of intact

soybean plants exposed to a variant pO^ have mechanisms that permit

maintenance of optimal internal pO^ for ATP production (Criswell e^ a2.,

1976). This kind of mechanism has not been demonstrated for Alnus,

however.

Substrate pH also affects nodulation and growth of European alder. The optimal pH of solution culture for the growth of nodulated

A. glutinosa lies between 4.2 and 5.4 (Ferguson, 1953) and is 5.4 for non-nodulated plants. Nodule formation occurs most freely over the pH range of 5.4 to 7.0 with the greatest number of nodules forming at pH

7.0 (Ferguson and Bond, 1953). Ferguson (1953) found that nodulation of A. glutinosa plants did not occur at a pH of 3.3 while 21

already-nodulated second year plants grew at pH 3.3. Best growth,

though, was achieved at pH 4.2. Tissues of A. glutinosa are well-

buffered with internal pH of 6.0 for the nodule and slightly below

5.0 for the root being optimal (Bond et , 1954). The capacity

of the host plant to tolerate relatively low pH levels seems to considerably exceed that of the nodule organism.

Plant growth and nitrogen fixation affect soil pH as well as being affected by it. Nyatsanga and Pierre (1973) believe that the effect of nitrogen fixation on soil acidity is greater than is gen­ erally recognized. Organic matter from the leaf and litter fall of nitrogen-fixing plants acts directly as a weak organic acid in soil. As.nitrogen is released from this decomposing organic matter, it is nitrified by soil micro-organisms resulting in greater acidity.

Nitrogen fixation also contributes more directly to soil acidity by generating 0.1 to 0.2 hydrogen ion per single nitrogen molecule assimilated (Raven and Smith, 1976). Hydrogen ions generated are excreted to the soil solution to counter intracellular acidity.

Ferguson (1953) observed a decrease of 0.3 to 0.4 pH units per day in nutrient solution containing A. glutinosa. Alder's ability to tolerate acid conditions makes it especially valuable for reforesting coal spoils (Lowry e£ , 1962).

Where nitrate-nitrogen is the only form of nitrogen available to A. glutinosa in solution culture, an alkaline pH drift results

(Quispel, 1958). Ammonium-nitrogen in solution culture under similar 22

conditions results in an acid drift. Species of combined nitrogen

interact with pH to affect alder growth. Ferguson (1953) demonstrated

that non-nodulated A. glutinosa supplied with sodium nitrate grew

well at all pH levels and was less sensitive to pH levels than plants

supplied with ammonium sulfate.

Under certain conditions, combined nitrogen exerts a harmful

effect on the nodulation of A. glutinosa. As levels of ammonium-

nitrogen are increased to 50 and 100 mg per 1, nodule development is

continually depressed (MacConnell and Bond, 1957). The presence of

any nitrate inhibits nodule formation as well (Quispel, 1958; Benecke,

1970). Pizelle (1965) found that the inhibitory effect of nitrate was not local, since nitrate supplied to a non-nodulated part of a Euro­ pean alder's root system suppressed nodule formation on the rest of the root system in a separate compartment not supplied with combined nitrogen. Wilson (1940) attributed the harmful effect of combined nitrogen on the nodule organism to two possibilities: (1) That a plant supplied with combined nitrogen, being more vigorous, is more immune to invasion by the nodule organism; and (2) that the early availability of nitrogen makes possible a more rapid synthesis of protein in the plant, thus lowering the level of carbohydrate in the tissues as compared with a plant in nitrogen-free solution.

Under other conditions, combined nitrogen is beneficial to the nodule organism of A. glutinosa. Low levels of ammonium-nitrogen result in increased plant and nodule development (Bond et al., 1954).

Nitrogen fixation per plant is considerably enhanced in the presence 23

of a low level of ammonium-nitrogen, owing to greater nodule development

(Stewart and Bond, 1961). Nodules are fewer but larger than those in

nitrogen-free solutions. Even at higher ammonium levels in excess of

plant needs, Stewart and Bond (1961) found that fixation per plant was

still comparable to that in nitrogen-free solution, though fixation per

unit weight of nodule tissue was somewhat lower than in nitrogen-free

solution.

Growth response of alder depends on the form of available nitrogen.

Ferguson (1953) determined that ammonium sulfate produced better growth

in non-nodulated A. glutinosa than sodium nitrate. Benecke (1970)

showed that maximum growth of nodulated A. viridis occurred when the

plants were grown independently of supplied nitrate-nitrogen. Nodulated

plants grew better in the absence of nitrate-nitrogen than non-nodulated

plants at any level of added nitrate-nitrogen. These results suggest

a saving of energy in the metabolism of ammonia or ammonium versus the

metabolism of nitrate.

Water availability is another limiting factor in nodulation and

nitrogen fixation. Sprent (1972) found that moisture stress in

the rooting medium of a legume caused a reduction in nitrogen fixation.

Reduced nitrogen fixation in nodulated plants under water stress

might be due to increased resistance to oxygen diffusion in the

membrane of stressed nodules (Parkhurst and Sprent, 1976) and the inhibition of shoot photosynthesis (Huang et al., 1975). Waterlogging can, however, be more severe than water stress in its effects on plant growth and nitrogen fixation (Minchin and Pate, 1975). 24

Substances produced by plants in competition with nitrogen-fixing

plants, such as gallic or tannic acid, can limit the establishment

and nodulation of nitrogen fixers (Blum and Rice, 1969).

Temperature has a pronounced effect on nitrogen fixation of A.

glutinosa. Wheeler (1971) determined that acetylene reduction reached a maximum at about 25°C which was six times faster than at 15°C and only slightly less at 30°C than at 25°C. Waughman (1972) found that the rate of acetylene reduction by detached nodules was positively correlated with temperature between 4 and 20°C, with the rate of reaction lower at 30°C than at 20°C. Optimal fixation of nitrogen by A. viridis nodules was shown by Benecke (1970) to occur at 20 C.

Low light levels and shortened daylength inhibit nodule formation in A. glutinosa (Quispel, 1954, 1958). The rate of acetylene reduction or nitrogen fixation by root nodules of young A. glutinosa plants increases substantially with increasing light intensity and declines rapidly when plants are kept in the dark (Wheeler and Bowes, 1974).

Maximal rates of nitrogen fixation in the root nodules of first- year A. glutinosa plants growing under natural illumination at nearly constant temperature were found by Wheeler (1969) to be attained at or after midday. Oxygen uptake and rate of ingress of photosynthate were also at a maximum around midday.

There is an overnight rise in total carbohydrate in A, glutinosa nodules suggesting that a substantial part of nodule carbohydrate is unavailable for nitrogen fixation and that maximal rates of fixation 25

occur only when new photosynthate is entering the nodules in quantity

(Wheeler, 1971). The midday peak in nitrogen fixation is not autonomous,

since after 24 hours in darkness, fixation falls to an insignificant

level (Wheeler, 1971), accompanied by a fall in nodule sugars,

notably sucrose.

Seasonal changes in nitrogen fixation and accumulation occur in

addition to diurnal changes. Stewart (1962) observed that during first season growth of A. glutinosa nitrogen fixation reached a maximum in late August but fell rapidly with the onset of autumn. Young nodules had the greatest fixation per unit of dry weight. Kirchner _et al. (1908), as cited in McVean (1955a), found that the nitrogen contents of roots,•shoots and bark doubles between the end of February and the end of May, after which there is a steady decline. 26

OBJECTIVES

The potential of alder as a silvicultural crop on nitrogen-poor

sites might be improved significantly by photosynthetic enhancement

of nitrogen fixation. A. rubra in the northwestern United States

competes favorably with associated species because of its photo-

synthetic efficiency coupled with its ability to symbiotically fix

nitrogen. Red alder has greater foliage surface per unit foliage

weight, greater total foliage area per seedling, and higher photo-

synthetic rate at high light intensities than associated coniferous

species (Krueger and Ruth, 1969).

Experiments in crop physiology indicate that increases in total

photosynthate available to plant nodules without decreases in photo-

synthate available to other tissue will result in increased nitrogen

fixation and growth (Havelka and Hardy, 1974; Hardy and Havelka, 1976).

The respiratory breakdown of photosynthate yields ATP energy,

reducing power and carbon skeletons that are used within the nodules

of alder for nitrogen fixation (Wheeler, 1969). Photosynthate demands

for nitrogen fixation may be similar to, or only slightly greater than,

requirements for the assimilation of combined nitrogen (Gibson, 1966;

Bergersen, 1971b; Minchin and Pate, 1973).

If selections of alder are to be made on the basis of photosynthetic

properties, with an eye towards the enhancement of nitrogen fixation, then the relationship between photosynthesis and nitrogen fixation for 27

specific combinations of host plant, endophyte and environment needs

to be studied.

Work of Gordon and Wheeler (1978) showed that 12 European alder clones differed significantly in rate of net photosynthesis per plant and rate of acetylene reduction per plant. Values for photosynthesis and acetylene reduction were significantly correlated for the 12 clones

(r = .59), suggesting that, barring differences in endophyte effective­ ness, selection of clones for high photosynthetic capability would usually result in the selection of clones with a high nitrogen fixation capacity.

The objectives of this study were to determine additional relationships between indicators of nitrogen fixation and indicators of photosynthesis and to determine the effect of host genotype on nitrogen fixation and the assimilation of combined nitrogen during early growth. 28

PROCEDURES

Plant Material

A. glutinosa seed was obtained from eatablished plantations in

Iowa and Illinois. The seed was sown in Jiffy peat pellets and grown under the same general cultural and environmental conditions in a greenhouse to obtain plant genotypes with a wide range of growth capacities (Bajuk et al., 1978). Selected plants were propagated by tip cuttings, and resulting clonal lines had a high probability of exhibiting genetically-based differences in growth capacities in a favorable, greenhouse environment.

Combined nitrogen was made available for plant growth during the selection process. Plants were fertilized regularly with a solution that included 144 ppm ammonium-nitrogen and 56 ppm nitrate-nitrogen.

Only one in 1000 plants formed nodules in uninoculated peat pellets.

Ten clonal lines were selected for use in this study (Table 1).

Several ramets were taken from one plant in each selected clonal line.

Plants from which ramets were taken were of the same age and had been grown under the same conditions. The basal end of each ramet was dipped in a 6000 ppm solution of indolbutyric acid in 50 percent ethanol for five seconds to promote rooting and was set in sterilized

Perlite. Plants were intermittently misted and kept in half-shade for eight weeks, by which time rooting had occurred in all clonal lines.

Misting and shading were gradually eliminated over the next three weeks. 29

Table 1. Growth of selected Alnus glutinosa cuttings under uniform conditions (Ba.juk et al., 1978)

Seed source of clonally- Clonal stock Mean total dry weight propagated plants number (g) after 6 weeks

Iowa Conservation Commission Nursery, Ames, Iowa 1-26 54.9

Iowa Conservation Commission Nursery, Ames, Iowa 2-50 49.1

4-H Camp, Madrid, Iowa 3-21 45.9

Iowa Conservation Commission Nursery, Ames, Iowa 1-23 44.5

Southern Illinois University Carbondale, Illinois 5-50 38.6

4-H Camp,.Madrid, Iowa 3-13 34.3

Carbondale, Illinois 4—40 27.6

Southern Illinois University Carbondale, Illinois 6-15 22.0

Southern Illinois University Carbondale, Illinois 5-40 10.0

Southern Illinois University Carbondale, Illinois 5-37 5.2

Plants were fertilized three times per week for three weeks to compensate for nutrients leached under mist and to establish healthy root systems. Peters 20-20-20 water-soluble fertilizer was applied with a Hydrocare proportioner to yield a solution containing 144 ppm ammonium-nitrogen, 56 ppm nitrate-nitrogen, 88 ppm phosphorous as available phosphoric acid, and 166 ppm potassium as potash. 30

Fertilization was then discontinued for two weeks, after which 60

alders were weighted and randomly assigned to treatments and positions

on benches in the same greenhouse bay.

Twenty alders, two from each of the 10 selected clonal lines,

were inoculated with 25 ml of a 10 percent, crushed-nodule preparation

(page 32) in 20 1 vessels of nitrogen-free Crone's solution (Table 2).

Two plants from each clonal line were put in 20 1 hydroponics-vessels

without crushed-nodule inoculum. Twenty more alders, two from each

clonal line, were potted in a mix consisting of equal parts, by volume, 3 of peat, vermiculite and Perlite with 9 kg per m of incorporated

MagAmp, a slow-release fertilizer.

In order to know the amount of total nitrogen and dry weight

accumulated during treatment, it was necessary to know initial dry weights and total nitrogen contents of experimental plants. Since these are destructive measurements, initial values in the experimental plants had to be estimated.

Eighteen alders remaining after assignment of plants to treatment vessels were weighed in the fresh state, dried in a 70°C oven for 48 hours, reweighed and analyzed for total nitrogen, including nitrate- nitrogen, using method B of Nelson and Sommer s (1973). Data are listed in the Appendix (Table A-20).

Regression equations to predict initial dry-weight from initial fresh-weight and initial total-nitrogen from initial fresh-weight were derived from this data: 31

GRAMS DRY WEIGHT = .14 GRAMS FRESH WEIGHT + .11 (r = .97).

GRAMS TOTAL N = .0029 GRAMS FRESH WEIGHT + .0043 (r = .97).

Estimates of initial dry-weights and initial total-nitrogen

contents of experimental plants listed in the Appendix are based on

these equations.

Plants were kept in treatment vessels from November 11, 1976 to

February 4, 1977. Continous hygrothermograph-recordings during this

period show a mean day-temperature in the greenhouse bay of 27°C

with a mean daily maximum of 33°C. Mean night temperature was 24°C

with a mean nightly minimum of 22°C. Relative humidity remained

stable at approximately 18 percent.

Supplemental lighting of low intensity was used to extend the

photoperiod to 16 hours. A combination of 200-watt, tungsten-filament,

clear-incandescent and 40-watt, cool-white, Ilucrescent lamps yielded 2 12 micro-Einsteins/m /sec. of physiologically active irradiance (PAI) at plant midcrowns. A Lambda LI 185 Quantum Radiometer with a pyran- ometer shielded to register energy between 400 and 700 nanometers in wavelength was employed for PAI measurements.

Nutrient Culture System

Each plant was grown in an individual hydroponics-unit consisting of a 20-1 plastic container with silver reflective exterior paint to prevent excessive heat-absorption. Each vessel was sealed with a wooden top containing a corked hole for adding water lost by transpiration and evaporation, a notch for aeration-tube insertion and a center hole 32

6 cm in diameter for the plant. A foam rubber strip around the base

of each alder stem inserted in a split, one-hole cork held the plants

upright in the central hole of the wooden top.

The vessels contained 20 1 of half-strength, nitrogen-free, Van

der Crone's solution (Table 2) with 1 ml/1 of modified Hoagland's

micronutrient supplement (Templeman, 1941).

Marine-aquarium air-stones, connected to an air compressor via

rubber tubing, copper pipe and flow regulators, provided a stream

of fine air bubbles in the nutrient solution.

Table 2. Nitrogen-free Van der Crone's solution (half-strength)

Nutrient Grams/liter

KCl .375

CaSO, ' .25 4 2H2O

MgSO^ • THgO .25

Fe^ (PO^ig • SHgO .125

Ca, (PO^), .125

Nodules used to prepare inoculum in this experiment originated

from 100 g of nodules that had been collected from the root system

of one European alder. This alder had been grown from seed at the Iowa

State Conservation Commission Nursery in Ames. The 100 g of original nodules were used to inoculate stock plants in nitrogen-free. Crone's solution. Nodules were propagated by reinoculation of additional stock plants. 33

A random sample of nodules from six different stock plants all

contained the spore-negative form of endophyte as determined by

Maurice Lalonde of the Kettering Research Laboratory in Yellow Springs,

Ohio. This suggests uniformity of endophytic material in at least

one characteristic.

Sixty grams of stock nodules were surface-sterilized by shaking

in a 10 percent clorox solution for five minutes followed by five one-minute rinses in distilled water. These nodules were ground with a mortar and pestle and diluted to a 10 percent by fresh weight solution in distilled water. Twenty-five milliliters of this single inoculum preparation were immediately added to vessels designated for this treatment. The inoculum was poured directly onto alder roots over the nutrient solution. The roots were then wholly submerged in the nutrient solution.

Added inoculum corresponded to an amount that caused formation of two-thirds the maximum number of nodules obtainable by Quispel

(1954) with European A. glutinosa nodules in 300 ml vessels without aeration.

Initial pH's of the unbuffered nutrient solutions at the time of inoculation ranged from 6.0 to 6.7. They were measured directly with a glass-electrode pH meter.

Alders grown in potting soil were fertilized three times per week with Peters fertilizer in the method previously described. 34

Acetylene-Reduction Assay

After discovery of the acetylene-reduction assay (Dilworth, 1966;

Schollhorn and Burris, 1966), it was shown to be a valuable index of nitrogen-fixing activity in both laboratory and field studies (Koch and Evans, 1966; Stewart ^ al., 1967; Hardy , 1968).

Acetylene acts as a noncompetitive inhibitor of nitrogenase, substituting for molecular nitrogen and consequently becoming reduced

(Hwang, Chen, and Burris, 1973). The ethylene produced can easily be detected and measured by gas chromatography. At volumes of 10 to 20 percent acetylene in air, there is no measurable competition with dinitrogen for the active site of nitrogenase in alder nodules

(Akkermans, 1971; Fleschner e^ £l., 1976).

Ferredoxin (Na^SjO^) is implicated in the reduction of the iron protein of nitrogenase (Burris, 1974). When the reduced iron protein binds MgATP, its potential is lowered to about -490 mv and it acquires the unique capacity to reduce the molybdenum-iron protein of nitrogenase.

The reduced molybdenum-iron protein probably serves as an electron sink for the reduction of dinitrogen and other substrates of nitrogenase including acetylene, reoxidizing the molybdenum-iron protein in the process.

On the basis of electron requirements, the production of three moles of ethylene from acetylene corresponds to the reduction of one mole of dinitrogen. The mean values for nitrogen fixation obtained from excavated nodules of A. tenuifolia and soybean by the 35

acetylene-reduction method agree with those obtained by the isotopic

^^2 technique (Fleschner et , 1976; Bergersen, 1970). Ratios of acetylene reduced to nitrogen fixed as determined by the method are close to the theoretical value of three.

All organisms known to fix nitrogen that have been tested by the acetylene-reduction procedure have produced ethylene stoichio- metrically (Becking, 1971). Although rigid standardization of the assay will be necessary before reliable, quantitative estimates can be derived, the acetylene-reduction assay is suitable for comparisons cf nitrogen-fixing systems or for studying the effects of imposed conditions within experiments (Bergersen, 1970). The acetylene- reduction assay was employed in this work in order to chart the development of nitrogenase activity during early growth of inoculated alders and to relate nitrogenase activity with leaf area development.

Whole-root-system assays avoid pitfalls associated with detached- nodule assays. Silver and Hague (1970), working with Myrica cerifera, found that young, non-woody, nodule tissue was more active in reducing acetylene than older woody tissue. The amounts of acetylene reduced by nodules of alder, even when the nodules are of similar age and taken from the same tree, may vary greatly (Waugham, 1972). In vivo assays for acetylene reduction by intact, attached root-systems also have an advantage in that repeated measurements of the same plant are possible.

Detectable levels of acetylene-reduction were generally established in the inoculated alders after three weeks. At this time, simultaneous 36

sampling of acetylene-reduction rates and leaf areas were begun. One

inoculated alder and an uninoculated control of the same clone were

measured per weekday (Monday through Friday). An inoculated and an

uninoculated plant from each of the 10 selected clones were measured

during the first two weeks in random order. During the next two

weeks the remaining alders in hydroponics were measured in the same

way, again in random order. Measurements of individual plants were

repeated after 28 days and a few plants were measured a third time,

56 days after initial measurements.

The lengths of leaves at midvein and their maximum widths were

measured. Leaf areas were estimated using a regression equation based

on the relationship between leaf area, determined with a planimeter,

and leaf length x leaf width:

LEAF AREA (cm^) = .78 (cm LENGTH x cm WIDTH) + .37 (r = .996)

The equation was derived from a random sample of 35 leaves gathered from stock plants (Table A-21). Leaves from all 10 clonal lines used in this study were included in the sample.

There were no significant differences in photosynthesis per unit leaf area among 12 clonal lines of European alder studied by Gordon and Wheeler (1978). Since eight of these clonal lines were used in this study, leaf area can probably be considered as a reasonable index of photosynthetic ability of these clones.

For the acetylene-reduction assay, attached root systems of whole plants were sealed in Saran Maraflex Shrink Bags which were obtained from American Can Company, American Lane, Greenwich, Connecticut. 37

The base of each stem and the middle of a short, L-shaped glass tube

were embedded in plasticine. A nylon cord, tied around the mouth of

the bag so as to press it tightly against the plasticine, effectively

sealed the root system in the saran bag. The flared outside end of

the glass tube was fitted with a serum stopper through which the bag

containing the root system was evacuated with a 50-cc syringe.

Fifty cubic centimeters of pure acetylene and 200 cc of air were

injected into each saran bag and each root system was incubated for

one hour at 25°C + 1°C. After incubation, four 1-cc samples were

withdrawn directly through the wall of each saran bag. Needles of

syringes were stuck in a rubber stopper to prevent escape of gaseous

sample and immediately transported from the greenhouse to a nearby

laboratory for detection of acetylene reduced to ethylene.

Samples were injected into the ports of a Varian model 2740 gas

chromatograph equipped with two columns and flame ionization detectors.

Columns were of stainless steel, 2.13 m long by 7.35 mm outside

diameter and were packed with Porapak R. Flow rates were 40 ml/min for the nitrogen carrier gas, 40 ml/min for hydrogen, and 400 ml/min for air. Column oven temperature was set at 60°C and detector tem­ perature at 110°C.

Samples of hydroponic media, uninoculated root systems, and air, all incubated with 20 percent acetylene, yielded no detectable ethylene.

Inoculated and uninoculated root systems, incubated without acetylene, produced no detectable ethylene. 38

Peak nitrogenase activity occurred at midafternoon, several

hours after peak solar radiation (Figure 1). All plants were

incubated during the late afternoon near sunset. By this time,

nitrogenase activity had returned from its peak to a steady basal

rate. Late afternoon incubation was to minimize day to day sampling

error that diurnal fluctuation in plant nitrogenase activity could

cause.

Photosynthesis

After completion of leaf area and acetylene reduction measurements, terminal leaf areas, photosynthesis rates and dark-respiration rates of the inoculated alders were measured.

Rates of photosynthesis were measured on whole plants in a plexiglass controlled environment chamber described by Broerman,

Gatherum and Gordon (1967). CO^ concentration was measured in a closed circuit system which included, in series, these elements:

1. A plexiglass, controlled environment chamber into which an

alder was sealed.

2. A spherical-float flowmeter with a flow-regulator valve.

3. A calcium-chloride desiccator.

4. An air filter.

5. A reference cell of a Beckman IR215 infra-red CO^ analyzer.

6. A rotary air pump pulling air through the infra-red analyzer. 39

O LU LO

LO 75 500 • I— on 5 a. Leaf Area ce: 2914 C/Vl O ZD O 60 400 si I oo LU LU O

O o CXl 45 300 - 2400

0400 0800 1200 1600 2000 2400 HOUR

Figure 1. Diurnal fluctuation in acetylene reduction by whole, nodulated root systems of Alnus glutinosa 40

Air flow was ^cintained at 2 1/min. Irradiance at plant 2 midcrown in the chamber was maintained at 2150 micro-Einsteins/m /sec

PAI as measured by a Lambda LI 185 Quantum Radiometer with pyranometer

shielded to detect radiation between 400 and 700 nanometers in

wavelength. Chamber temperature was constant at 24°C.

Each alder, with its roots in a 2-1 beaker of nutrient solution,

was preconditioned in the chamber for 45 minutes. The chamber was

sealed and net photosynthesis was calculated as the tangential rate

of CO^ uptake at CO^ concentration of 320 ppm in the chamber. The

chamber was then darkened and dark respiration was monitored for 60

minutes. A stable, dark-respiration rate became established several

minutes after photosynthesis stopped and was recorded.

Tissue Analysis

Eighty-five days after assignment to treatment vessels, final pH measurements of the nutrient solutions were made and the alders were harvested. The leaves, stems, roots and nodules of each alder were placed in separate paper bags for drying. Counts were made of the number of nodules per plant. Only nodules that were at least 2 mm across at their widest point were counted. All plant tissue was dried in a 70°C oven for 48 hours and weighed. Tissues were ground in a

Wiley mill to pass through a 40-mesh screen and redried for 24 hours.

Kjeldahl determination of total nitrogen, including nitrate-nitrogen, in the plant tissues was performed following procedure B of Nelson and

Sommers (1973). 41

All data are listed in the Appendix.

Since the number of experimental plants was limited by available space and equipment, treatments were replicated only twice in order to include the maximum number of alder clones.

Quantitative information on environmental gradients within the greenhouse bay, which would have been the basis for blocking, was lacking, so a completely randomized experimental design was employed.

Statistical methods used in this work include multiple regression, simple linear regression, analysis of variance, correlation and con­ fidence intervals (Cochran and Cox, 1957; Snedecor and Cochran, 1967). 42

RESULTS

Reasonable care was taken at the beginning of this experiment to

obtain rooted cuttings of uniform sizes at the time of assignment to

treatments, initial dry weight estimates of rooted cuttings ranged

from .59 to 1.53 g with corresponding initial nitrogen estimates from

14 to 34 mg/plant. There were no significant differences among clones

in these initial values.

Multiple regression analysis of leaf area (Table 3) and acetylene

reduction (Table 4) shows that little variation can be associated with

differences in clonal lines. Data (Tables A-2 and A-3) were collected

after establishment of acetylene reduction in inoculated plants in a

repeated, random order over a two month period. Each of 20 plants

was sampled at least twice, although two of the 20 inoculated plants

that did not form nodules were deleted from these analyses.

Nitrogenase activity, leaf expansion and early growth of alder

in nitrogen-free culture depend on successful establishment of the

nitrogen-fixing symbiont. Even though all plants were rooted under

closely similar conditions and the same amount of a single inoculum

preparation was applied at the same time to all plants, inconsistent

nodulation occurred. Of 20 plants that were inoculated, IS formed

nodules. The appearance of visible nodules on roots acquired from

10 to 28 days. 43

Table 3. Analysis of variance from regression on dependent variable leaf area

Source of variation Degrees of Sequential F-value Probability freedom mean squares level

Linear regression 850,414 20.6012 ,0005 on time

Quadratic regression 209,025 5.0636 .0369 on time

Clones 9 80,644 1.0413 n.s.

Plants within clones 8 77, 744

Residual 16 41,280

Table 4. Analysis of variance from regression on dependent variable nitrogenase activity

Source of variation Degrees of Sequential F-value Probability freedom mean squares level

Linear regression 1 299,655,172 15.9212 .0013 on time

Quadratic regression 1 17,686,172 0.9397 .6513 on time

Clones 9 22,899,603 0.5855 n.s.

Plants within clones 8 39,109,341

Residual 16 18,821,181 44

Based on the Quispel curve (1954), the amount of inoculum used in this study should have produced two-thirds of the maximum number of nodules obtainable. However, nodulation was inconsistent (Table

A-7), suggesting that the North American inoculum used in this work was less infective than the European inoculum used by Quispel (1954b).

A spore or bacteroid form of the alder endophyte is thought to

be an infective agent (Akkermans and Van Dijk, 1976). These bacteria-

like cells have not been described in plant species that nodulate with

difficulty in water culture, such as Coriaria spp., while plant species

that can be readily nodulated in water culture with suspensions of

their respective crushed root nodules, such as Alnus spp., Hippophaë

rhamnoides, Eleagnus spp. and Gale palustris, have been reported to

harbor endophytes that produce bacteria-like cells (Becking, 1970b).

The occurrence of an endophytic type that does not form spores or bacteria-like cells (spore form) was noted by Schaede (1933) and Van

Dijk and Merkus (1976) in Alnus glutinosa. Maurice Lalonde of the

Kettering Research Laboratory in Yellow Springs, Ohio, found only the spore form of endophyte in a sample of nodules from six stock plants from which nodules were gathered to prepare inoculum for this work.

If these spores are the chief agent of root infection in Alnus, then fewer numbers of infective i .rticles associated with the spore type would have reduced the chance of contact between root hairs and 45

infective particles, resulting in fewer nodules and greater periods

of time required for infection to occur. Difficulties in establishment

could also be attributed to lesser affinity between A. glutinosa and an

unusual North American endophyte. Lack of uniformity in the time of infection and the establishment

of nitrogen fixation in nodules resulting from low density of infective

particles might have masked real differences attributable to host

genotype during early growth.

Relationships between indicators of nitrogen fixation and

indicators of photosynthesis established across host genotypes are

strong. Regression of late-afternoon nitrogenase activity on total

leaf area of individual plants is illustrated in Figure 2. Each of

the 20 inoculated plants contributes two or three points to the figure.

Measurements of the growing plants were made in random order and

repeated at 28-day intervals (Tables A-2 and A-3). Figure 3 represents

the regression of photosynthesis rate, under uniform conditions, on

total nitrogen content of inoculated alders grown in nitrogen-free water culture. Measurements of photosynthesis rates were made on all 20 inoculated plants at the same time, just prior to harvest (Table A-5).

Nitrogen contents were derived from subsequent Kjeldahl-nitrogen analysis of dried tissue (Table A-9). In both cases correlation coefficients are significant at the 1 percent level.

At time of harvest, the fertilized alders had accumulated the greatest amount of nitrogen (Table 5). 46

Table 5. Total plant nitrogen (mg) at harvest

Treatment Number of Mean and 95% Range observations confidence interval

Nodulated 18 245.8 + 97.1 26.7 - 661.9

Fertilized 20 862.9 + 243.9 137.9 -1822.1

No nitrogen 20 20.0+ 2.5 9.1- 30.3 47

75

r=.94 60

45 •V.

2 30

15

0 1000 2000 3000 LEAF AREA-CM?

Figure 2. Regression of late-afternoon acetylene reduction rate on leaf area of nodulated Alnus glutinosa 48

200

5 = 15. r=.89 ZD O •I Csj O

O S CO

Q_ I I I I 1 • I L_ 100 200 300 400 500 600 700 NITROGEN-MG/PIANT

Figure 3. Regression of photosynthesis rate on total-plant nitrogen of nodulated Alnus glutinosa at time of harvest 49

Nitrogen content of nodulated alders was considerably greater

than that of uninoculated alder controls grown in the same nitrogen-

free substrate.

Percentages of nitrogen in alder on a dry-weight basis were

greatest in the leaf tissue of all treatment groups, followed by

percentages of nitrogen in nodules, roots, and lastly, stems (Table 6).

Table 6. Percentage nitrogen in alder tissue (mean values and 95 percent confidence intervals) at harvest

Treatment Number of Whole Leaves Stem Roots Nodules observations plant

Nodulated 18 2.71+.29 3.99+.29 1.55+.18 1.81+.10 3.68+.27

Fertilized 20 3.12+.19 4.52+.18 1.56+.19 2.33+.18

No nitrogen 20 1.19+.07 1.41+.16 0.68+.06 1.39+.10

Since detectable levels of nitrogenase activity could not be

generally established in the inoculated alders until three weeks

after inoculation, with nodules becoming visible from 10 to 28 days

after inoculation, it is not surprising that fertilized alders

accumulated greater amounts of nitrogen during the same early growth

period.

As nitrogen availability to alders increased, so did the ratio of shoot to root growth (Table 7). 50

Table 7. Shoot/root ratios at harvest

Treatment Number of Mean and 95% Range observations confidence interval

Inoculated 20 1.99 + .49 .45 - 3.81

Fertilized 20 4.86 + .32 4.05 - 6.45

No nitrogen 20 .69 + .08 .45 - .95

The development of more root tissue in relation to shoot tissue

would have some survival advantage for alder where nitrogen is limiting.

With greater relative root growth there would be a greater chance of

root-contact with infective particles of the actinomycetal endophyte

Significant differences in dry weight and nitrogen content of

alders occurred among alder clonal lines grown on combined nitrogen

(Table 8). Nodulated alders and controls without nodules grown in

nitrogen free water culture did not exhibit significant differences

among clonal lines (Tables 9 and 10). Since infection of inoculated

alders, as indicated by appearance of nodules and the occurrence of

detectable levels of acetylene reduction, was not uniform in time or

quantity, it is not surprising that any effect that host-plant geno­

type might have had on growth of inoculated plants was not evident.

In future work, efforts to determine the infectivity of different

endophytic inocula and to better control such plant characteristics

such as number of root hairs, leaf area, carbohydrate status, and nitrogen status at time of inoculation might result in better control 51

over the timing and quantity of infection. More research will be

necessary to develop such procedures.

In general, clonal lines with faster growth rates on combined

nitrogen had lower nitrogen percentages than slower-growing clones

(Figure 4) and the differences in nitrogen percentages among clones were significant (Table 8). This suggests that differences in growth among clones where nitrogen is not limiting were attributable to the plant's ability to use rather than to take up nitrogen. Among the inoculated alders, plants that had the lowest nitrogen percentages were poorly nodulated (Table A-7) and had achieved the lowest dry mass

(Table A-6), illustrating that nitrogen availability to the plant, as well as the plant's ability to use nitrogen, can limit alder growth.

Table 8. Mean squares from analysis of variance for results of experiments with fertilized alder clones

Source Degrees of Total plant Total plant Total plant freedom dry weight nitrogen nitrogen percent

Clone 9 637.34 .53 .30 (.0001*) (.0002) (.0172)

Error 10 42.32 .04 .07

*Denotes probability level.

Linear correlation coefficients for relationships between average clonal values of variables determined at time of harvest are illustrated in Table 11. 52

Table 9. Mean squares from analysis of variance for results of experiments with nodulated alder clones

Source Degrees of Total plant Total plant freedom dry weight nitrogen

Clone 9 41.25 (.4932*) .04 (.3979)

Error 8 40.37 .03

*Denotes probability level.

Table 10. Mean squares from analysis of variance for results of experiments with uninoculated alder clones

Source Degrees of Total plant Total plant freedom dry weight nitrogen

Clone 9 .23 (.4827*) .000027 (.5722)

Error 10 .22 .000031

*Denotes probability level. 60 • DRY MASS |0, 50

5 40 5; Ln CD 30 Ln 3> U> S CO m 20

m 10

2-50 1-23 1-26 4-40 3-21 3-13 5-50 6-15 5-40 5-37 ALDER CLONE

Figure 4. Clonal means for dry mass and percentage nitrogen of fertilized Alnus glutlnosa plants at time of harvest 54

There is no significant correlation between variables associated with growth of uninoculated alder clones in nitrogen-free solution and variables of the same clones grown on fixed or combined nitrogen.

However, total dry-weight and nitrogen content means of fertilized alder clones are significantly, positively correlated with means for achieved photosynthesis and respiration rates, leaf area, and nodule dry-weight of inoculated alder clones. Total dry-weight means of inoculated alder clones approach significant positive correlation with dry-weight means of fertilized clones. These correlations suggest that predictions of host-genotype performance with a symbiont might be based on growth performance, under similar environmental conditions, of the same host genotype on combined nitrogen.

The correlation matrix also reveals that, in every case, dry weight of nodule tissue is more highly correlated with values for growth and nitrogen fixation in the inoculated alder clones than is number of nodules. Mean clonal dry weight of nodule tissue is cor­ related with clonal means of the same nodulated plants for achieved photosynthesis and respiration rates, total dry weight, nitrogen content, leaf area and leaf nitrogen content (Table 11). Correlation coefficients for relationships between mean number of nodules per plant per clone and mean values for growth and nitrogen fixation in the inoculated alder clones are low and not significant. Dry weight of nodule tissue is a better indicator of symbiont effectiveness than is number of nodules. Gibson (1969) also suggested that nodule dry Table 11. Linear correlation coefficients and probability levels for relationshiops between mean clonal values of variables determined at time of harvest

Inoculated Variable Photo- Respiration Total Total Leaf area synthesis dry weight nitrogen

Photo­ 1.00 synthesis (.0000*) Respira­ .96 1.00 tion (.0001) (.0000) Total dry .93 .91 1.00 weight (.0001) (.0002) (.0000) Total .90 .88 .98 1.00 nitrogen (.0001) (.0008) (.0001J (.0000) Leaf .87 .88 .94 .88 1.00 area (.0009) (.0007) (.0001) (.0007) (.0000) Leaf .88 .85 .96 .997 .86 nitrogen (.0009) (.0019) (.0001) (.0001) (.0012) Nr. of .42 .45 .45 .31 .55 nodules (.2326) (.1909) (.1958) (.3S36) (.0989) Dry weight .95 .93 .95 .90 .97 of nodules (.0001) (.0001) (.0001) (.0004) (.0001) Change in -.77 -.77 -.74 -.63 .83 uH (.0088) (.0090) (.0150) (.0523) (.0029) Total dry .68 .73 .61 .46 .72 weight (.0303) (.0170) (.0596) (.1759) (.0190) Total . 66 .71 .61 .46 .71 nitrogen (.0371) (.0208) (.0622) (.1805) (.0225) Total dry .36 .47 .43 .43 .42 weight (.3056) (.1732) (.2139) (.2174) (.2290) Total .33 .38 .39 .44 .31 nitrogen (.3521) (.2345) (.2662) (.2040) (.3842) Change in -.21 -.24 -.01 .12 -.15 PH (.5587) (.5105) (.9786) (.7507) (.6739)

*Denotes probability level. 56

Inoculated Fertilized Leaf No. of Dry weight Change Total dry Total nitrogen nodules of nodules in pH weight nitrogen

1.00 (.0000) .25 1.00 (.4861) (.0000) .88 .53 1.00 (.0008) (.1154) (.0000) —.58 —.84 —.86 1.00 (.0813) (.0024) (.0016) (.0000) .41 .57 .70 -.72 1.00 (.2369) (.0878) (.0238) (.0185) (.0000) .41 .56 .68 -.69 .997 1.00 (.2414) (.0916) (.0301) (.0272) (.0001) (.0000) .40 .49 .38 -.49 .28 .28 (.2486) (.1507) (.2779) (.1496) (.4300) (.4325) .43 .30 .32 -.32 .08 .09 (.2138) (.4005) (.3628) (.3744) (.8309) (.8151) .17 -.40 -.18 .38 -.54 -.52 (.6447) (.2541) (.6118) (.2787) (.1079) (.1241) 57

Table 11. Continued

No nitrogen Variable Total dry- Total Change weight nitrogen in pH

Photo­ synthesis Respira­ tion Total dry weight Total nitrogen Leaf area Leaf nitrogen No. of nodules Dry weight of nodules Change in _PH Total dry weight Total nitrogen

Total dry 1.00 weight (.0000) Total .91 1.00 nitrogen (.0002) (.0000) Change in .31 .53 1.00 pH (.3870) (.1182) (.0000) 58

weight is the best available index of bacteroid tissue in assessing the effectiveness of nitrogen fixation in nodulated plants.

The mean clonal decrease in pH of the unbuffered nutrient solutions was correlated with clonal growth values and nitrogen contents of inoculated alders. Solution pH's were decreased by as much as 2.5 units during this experiment. The net effect of growing European alder symbiotically in nitrogen-free solution is an increase in hydrogen ion concentration. 59

DISCUSSION

Strong relationships exist between indicators of photosynthesis

and nitrogen fixation in European alder plants. Significant positive

correlations exist between plant leaf area and late-afternoon acetylene

reduction (.94), between photosynthesis rate and plant nitrogen content

(.89), and between photosynthesis rate and dry weight of nodule

tissue (.95) for European alders inoculated and grown in nitrogen-free

solution cultures.

Significant differences in growth and nitrogen content occur

among alder clones provided with a complete nutrient preparation

including combined nitrogen. However, growth differences attributable

to host-plant genotype of nodulated alders grown in nitrogen-free

Van der Crone's solution were not significant. Since the timing and

quantity of nodule formation varied considerably, it is not surprising

that no effect of host plant genotype on growth of inoculated alders

could be observed.

Inoculated plants were treated with equal portions of the same crushed-nodule preparation. However, the nodular endophyte seems to have been a North American strain present in Iowa soil that does not produce abundant spores or granule. A popular theory holds that these spores are infective particles of the alder endophyte (Van

Dijk and Merkus, 1976). The low infectivity of this crushed nodule preparation suggests an explanation for the inconsistent nodular development of inoculated plants. That is, the lack of spores 60

associated with the endophyte used in this work resulted in fewer

nodules and longer time periods required for the chance contact

between infective particles and root hairs to occur than experienced

by Quispel (1954) with the same quantity of European nodule

inoculum. Difficulties in establishment of nodules could also be

attributed to lesser affinity between European A. glutinosa and a

North American endophyte (Lalonde and Quispel, 1977).

Greater variation in the time of nodule formation was probably

more important than variation in the number of nodules formed in

covering any growth effects that might have been caused by host-plant

genotype. Dry weight of nodular tissue was better correlated by far

with measurements of growth and nitrogen assimilation than was nodule

number per plant.

Although growth differences among alder clones were not significant

for inoculated plants, results suggest that an alder genotype's ability

to develop a symbiotically-fixed nitrogen supply can be predicted by

its growth performance on combined nitrogen. Significant positive

correlation exists between dry mass production of alder clones on

combined nitrogen and dry mass production of nodules of the same clones grown in nitrogen-free solution (.70). Development of nodular

tissue mass might be an early indication of clonal differences in growth in a nitrogen-free environment. Nodules might receive priority in photosynthate allocation during this early growth period. Dry mass and nitrogen content of alder clones grov^ on combined nitrogen were not as well correlated with total dry mass of the same clones 61

inoculated and grown in nitrogen-free solution (both r values = .61)

as with nodular tissue weight. Other early indicators of the success­

ful establishment of a symbiotic relationship in alder clones that

were significantly, positively correlated with the same clones'

growth performance on combined nitrogen include achieved photosynthesis

and respiration rates and leaf area.

Differences in early growth of alder on combined nitrogen

compared with growth of alder establishing a symbiotic relationship were considerable. On the average, alders grown on combined nitrogen had three to four times the dry mass of alders inoculated and grown in nitrogen-free hydroponics. This difference should not be confused, however, with differences between the energetic efficiencies of nitrogen fixation and combined nitrogen assimilation. During the early stages of nodule initiation and development, there is a definite carbohydrate requirement that cannot be measured against nitrogen fixation, but which causes the nodulated plants to weigh significantly less than control plants (Gibson, 1966). Three weeks passed after inoculation of alders before even low levels of acetylene reduction were detected in this work. Relative growth rates of fertilized and nodulated plants may thereafter be similar.

There seemed to be a steady base rate of acetylene reduction in nodulated alders, upon which is superimposed activity stimulated by current photosynthetic products (Figure 1). Although there is an overnight rise in total alder-nodule carbohydrate, the maximum acetylene-reduction activity occurs with the increased influx of new 62

photosynthate after noon, suggesting that a substantial part of nodule

carbohydrate is unavailable for fixation (Wheeler, 1971). Pea plants also rapidly use new photosynthate for acetylene reduction (Lawrie and Wheeler, 1973).

Alders assayed in this study were larger than those used in previous studies (Wheeler, 1969) and the peak acetylene reduction period occurred later in the afternoon, presumably at a time of maximum influx of new photosynthate. The basal, endogenous rate of acetylene reduction was greater in larger plants. Soybean nodules also exhibit a basal, endogenous rate (Bergersen, 1970).

The potential expressed by alder or other plant genotypes under ideal conditions may not be realized under different conditions in the field (Wacek and Brill, 1976) or with different endophytes

(Brill, 1977). Although this work has revealed certain relationships and patterns that may be of use in increasing the efficiency of alder's nitrogen fixing symbiosis, it is important that the work be extended under both field and refined, laboratory conditions. Field plantings on sites where these clones might be useful, such as on coal spoils and in mixed plantations, should be installed at various locations in the central United States. Better techniques for obtaining more control over root infection need to be developed and used. Combinations of these alder hosts and additional endophytes need to be tested for symbiotic efficiency. Perhaps, then, benefits of increased produc­ tivity and soil improvement that these plants can provide will be realized. 63

ACKNOWLEDGMENTS

Special thanks are due to my wife Norine for her patience during the course of this work; to John Gordon for research support and guidance; to Paul Hinz for statistical help; and to Rich Faltonson for assistance in the propagation and care of experimental plants. 64

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APPENDIX 77

Table A-1. Data on inoculated alders in hydroponics

Clone Plant Initial Initial Initial dry weight total N nutrient estimate (g) estimate (g) solution pH

1-26 1 1.26 .0281 6.5 1-26 2 0.98 .0222 6.1 2-50 1 0.91 .0208 6.0 2-50 2 0.85 .0197 6.2 3-21 1 1.29 .0287 6.2 3-21 2 1.45 .0321 6.0 1-23 1 1.03 .0234 6.1 1-23 2 0.82 .0191 6.7 5-50 1 1.41 .0313 6.3 5-50 2 1.03 .0234 6.3 3-13 1 0.81 .0188 6.1 3-13 2 1.37 .0304 6.3 4-40 1 1.17 .0263 6.4 4-40 2 0.77 .0179 6.2 6-15 1 1.37 .0304 6.0 6-15 2 1.37 .0304 6.0 5-40 1 0.94 .0214 6.2 5-40 2 0.59 .0142 6.5 5-37 1 0.60 .0145 6.4 5-37 2 0.-7 .0159 6.4 78

Table A-2. Data on inoculated alders in hydroponics

Clone Plant Date of First First nitrogenase first sample leaf area activity (cm2) (n moles C H /hr/plant)

1-26 1 Dec. 6 168.2 355.0 1-26 2 Dec. 23 81.6 0.0 2-50 1 Dec. 13 212.7 2980.0 2-50 2 Dec. 31 228.7 3412.5 3-21 1 Dec. 17 113.0 7.5 3-21 2 Dec. 29 185.4 5212.5 1-23 1 Dec. 10 57.4 35.0 1-23 2 Dec. 20 216.9 3507.5 5-50 1 Dec. 16 129.5 37.5 5-50 2 Dec. 28 373.7 6465.0 3-13 1 Dec. 7 142.5 462.5 3-13 2 Dec. 24 110.4 55.0 4-40 1 Dec. 8 122.0 230.0 4-40 2 Dec. 30 91.5 2.5 6-15 1 Dec. 15 121.4 90.0 6-15 2 Dec. 21 113.2 132.5 5-40 1 Dec. 9 99.6 5.0 5-40 2 Dec. 27 78.7 247.5 5-37 1 Dec. 14 79.2 0.0 5-37 2 Dec. 22 193.0 3595.0 79

Table A-3. Data on inoculated alders in hydroponics

Clone Plant Date of Second Second nitrogenase second sample leaf area activity (cm^) (n moles C2H^/hr/plant)

1-25 1 Jan. 3 429.8 8160.0 1-26 2 Jan. 20 73.9 0.0 2-50 1 Jan. 10 978.7 4380,0 2-50 2 Jan. 28 921.7 18,697.1 3-21 1 Jan. 14 85.7 975.0 3-21 2 Jan. 26 529.7 6285.0 1-23 1 Jan. 7 93.1 1400.0 1-23 2 Jan. 17 709.9 23,842.5 5-50 1 13 197.3 2315.0 5-50 2 Jan. 25 1354.5 14,942.5 3-13 1 Jan. 4 447.7 14,737.5 3-13 2 Jan. 21 249.6 2067.5 4-40 1 Jan. 5 112.5 2129.4 4-40 2 Jan. 27 30.9 2.5 6-15 . 1 Jan. 12 206.9 1572.5 6-15 2 Jan. 18 160.3 3060.0 5-40 1 Jan. 6 57.6 295.3 5-40 2 Jan. 24 107.9 670.0 5-37 1 Jan. 11 28.2 373.2 5-37 2 Jan. 19 519.4 7340.0 80

Table A-4. Data on inoculated alders in hydroponics

Clone Plant Date of Third Third nitrogenase third sample leaf area activity (cm2) (n moles C^H^/hr/plant)

1-26 1 Jan. 31 2061.0 41,337.7 1-26 2 2-50 1 Feb. 1 2914.0 74,984.9 2-50 2 3-21 1 Feb. 2 906.7 29,355.3 3-21 2 1-23 1 1-23 2 5-50 1 5-50 2 3-13 1 3-13 2 4—40 1 4—40 2 6-15 1 6-15 2 5-40 1 5-40 2 5-37 1 5-37 2 81

Table A-5. Data on inoculated alders in hydroponics

Clone Plant Terminal Terminal Terminal nutrient leaf area photosynthesis solution pH (cm^) rate (mg CO^/hr/plant)

1-26 1 4.3 2061.0 136.76 1-26 2 6.0 39.1 1.61 2-50 1 4.2 2914.1 99.98 2-50 2 4.5 1598.2 80.18 3-21 1 6.1 238.6 18.61 3-21 2 5.0 906.7 67.22 1-23 1 5.4 190.5 16.66 1-23 2 4.2 2059.2 174.73 5-50 1 5.2 716.4 47.03 5-50 2 4.2 2565.6 115.16 3-13 1 4.3 1774.6 136.76 3-13 2 5.4 763.1 45.37 4-40 1 5.2 732.8 49.23 4-40 2 6.2 24.6 1.47 6-15 1 5.2 619.9 45.58 6-15 2 5.2 351.1 41.96 5-40 1 6.0 51.9 2.96 5-40 2 6.4 213.5 15.04 5-37 1 6.4 94.9 5.91 5-37 2 4.8 1249.0 75.72 82

Table A-6. Data on inoculated alders in hydroponics

Clone Plant Terminal dark Terminal Terminal respiration rate leaf stem (mg CO^/hr/plant) dry weight dry weight (g) (g)

1-26 1 22.23 9.9669 5.0019 1-26 2 .03 .2495 .4908 2-50 1 24.51 8.4868 6.134 2-50 2 10.15 4.4174 2.2384 3-21 1 6.62 1.0413 .4767 3-21 2 18.21 3.5380 1.5809 1-23 1 5.39 1.0193 .6696 1-23 2 26.16 7.9813 4.5649 5-50 1 8.28 2.6025 1.1443 5-50 2 24.12 9.4321 5.3906 •y 3-13 a. 19.48 6.5160 3.1713 3-13 2 14.45 2.3740 .8313 4-40 1 9.21 2.9561 1.1301 4-40 2 .46 .1325 .2949 6-15 1 10.05 2.5974 1.3304 6-15 2 7.11 2.0503 1.0679 5-40 1 1.74 .2921 .3788 5-40 2 5.61 .8239 .3667 5-37 1 .00 .3017 .3575 5-37 2 12.61 5.5950 2.0662 83

Table A-7. Data on inoculated alders in hydroponics

Clone Plant Terminal Terminal Terminal Terminal root nodule total number dry weight dry weight dry weight of (g) (g) (g) nodules

1-26 1 . 3.6629 .4912 19.1229 48 1-26 2 1.1583 .0000 1.8986 0 2-50 1 3.1828 .6552 18.4589 72 2-50 2 1.9632 .2304 8.8495 55 3-21 1 1.4542 .0685 3.0407 19 3-21 2 2.6094 .2113 7.9396 41 1-23 1 1.0466 .0739 2.8094 4 1-23 2 3.3283 .5797 16.4542 138 5-50 1 2.4879 .1499 6.3846 33 5-50 2 5.5008 .5302 20.8537 125 3-13 1 2.3937 .5304 12.6114 35 3-13 2 1.8727 .1442 5.2222 39 4-40 1 1.7887 .1640 6.0383 47 4-40 2 .9576 .0000 1.3850 0 6-15 1 1.9885 .1727 6.0890 28 6-15 2 1.6387 .1297 4.8866 17 5-40 1 .8015 .0100 1.4824 2 5-40 2 .6845 .0464 1.9215 32 5-37 1 .6381 .0338 1.3311 15 5-37 2 2.0824 .3680 9.7436 61 84

Table A-8. Data on inoculated alders in hydroponics

Clone Plant Terminal Terminal Terminal leaf N stem N root N (g) (g) (g)

1-26 1 .4565 .0755 .0813 1-26 2 .0035 .0032 .0169 2-50 1 .2818 .0626 .0665 2-50 2 .1979 .0405 .0318 3-21 1 .0397 .0062 .0253 3-21 2 .1500 .0289 .0445 1-23 1 .0318 .0078 .0179 1-23 2 .2171 .0758 .0686 5-50 1 .1119 .0191 .0388 5-50 2 .4612 .0825 .0979 3-13 1 .2913 .0553 .0452 3-13 2 .0964 .0205 .0296 4-40 1 .1230 .0188 .0315 4-40 2 .0023 .0026 .0128 6-15 1 .1140 .0193 .0314 6-15 2 .0851 .0133 .0279 5-40 1 .0851 .0039 .0133 5-40 2 .0330 .0054 .0125 5-37 1 .0114 .0046 .0124 5-37 2 .2350 .0415 .0458 85

Table A-9. Data on inoculated alders in hydroponics

Clone Plant Terminal Terminal nodule N total N (g) (g)

1-26 1 .0197 .6330 1-26 2 .0000 .0236 2-50 1 .0231 .4340 2-50 2 .0103 .2805 3-21 1 .0021 .0733 3-21 2 .0072 .2310 1-23 1 .0023 .0598 1-23 2 .0217 .3832 5-50 1 .0058 .1756 5-50 2 .0203 .6619 3-13 1 .0208 .4126 3-13 2 .0063 .1528 4-40 1 .0066 .1799 4-40 2 .0000 .0177 6-15 1 .0066 .1713 6-15 2 .0053 .1316 5-40 1 .0004 .0267 5-40 2 .0013 .0522 5-37 1 .0008 .0292 5-37 2 .0139 .3362 86

Table A-10. Data on uninoculated alders in hydroponics

Clone Plant Initial Initial Initial dry weight total N nutrient estimate (g) estimate (g) solution pH

1-26 ; 1 .75 .0176 6.2 1-26 : 2 1.09 .0246 6.4 2-50 ; 1 .85 .0197 6.2 2-50 2 .85 .0197 6.3 3-21 1 .84 .0194 6.5 3-21 2 1.30 .0290 6.1 1-23 1 .77 .0179 6.2 1-23 2 .83 .0191 6.6 5-50 1 .88 .0203 6.1 5-50 2 1.24 .0278 6.3 3-13 1 .89 .0205 6.2 3-13 2 .68 .0162 6.5 4-40 1 .96 .0220 6.2 4-40 2 .81 .0188 6.4 6-15 1 .91 .0208 6.2 6-15 2 .75 .0176 6.2 5-40 1 .53 .0130 6.6 5-40 2 .75 .0176 6.4 5-37 1 1.10 .0249 6.0 5-37 2 .67 .0159 6.2 87

Table A-11. Data on uninoculated alders in hydroponics

Clone Plant Date of First First nitrogenase first sample leaf area activity (cm^) (n moles C^H^/hr/plant)

1-26 1 Dec. 6 79.8 1-26 2 Dec. 23 143.0 2-50 1 Dec. 13 137.1 2-5C 2 Dec. 31 78.4 3-21 1 Dec. 17 76.0 3-21 2 Dec. 29 77.8 1-23 1 Dec. 10 115.6 1-23 2 Dec. 20 115.4 5-50 1 Dec. 16 99.9

5-50 2 Dec. 28 156.4 — 3-13 1 Dec. 7 110.8 3-13 2 Dec. 24 91.2 4—40 1 Dec. 8 118.1 4-40 2 Dec. 30 83.8 6-15 1 Dec. 15 111.4 6-15 • 2 Dec. 21 38.4 5-40 1 Dec. 9 53.3 5-40 2 Dec. 27 86.7 5-37 1 Dec. 14 94.8 — 5-37 2 Dec. 22 125.5 88

Table A-12. Data on uninoculated alders in hydroponics

Clone Plant Date of Second Second nitrogenase second sample leaf area activity (cm2) (n moles C^H^/hr/plant)

1-26 1 Jan. 3 43.7 1-26 2 Jan. 20 79.3 2-50 1 Jan. 10 151.1 2-50 2 Jan. 28 38.2 3-21 1 Jan. 14 6518 3-21 2 Jan. 26 51.2 1-23 1 Jan. 7 136.3 1-23 2 Jan. 17 55.3 5-50 1 Jan. 13 86.8 5-50 2 Jan. 25 121.3 3-13 1 Jan. 4 114.2 3-13 2 Jan. 21 61.5 4—40 1 Jan. 5 66.1 4—40 2 Jan. 27 46.5 6-15 1 Jan. 12 73.7 6-15 2 Jan. 18 28.6 5-40 1 Jan. 6 41.6 5-40 2 Jan. 24 0.0 5-37 1 Jan. 11 15.7 5-37 2 Jan. 19 35.8 89

Table A-13. Data on uninoculated alders in hydroponics

Clone Plant Date of Third Third nitrogenase third sample leaf area activity (cm^) (n moles C2H^/hr/plant)

1-26 1 Jan. 31 36.3 1-26 2 2-50 1 Feb. 1 142.7 2-50 2 3-21 1 Feb- 2 51.4 3-21 2 1-23 1 1-23 2 5-50 1 5-50 2 3-13 1 3-13 2 4-40 1 4-40 2 6-15 1 6-15 2 5-40 1 5-40 2 5-37 1 5-37 2 90

Table A-14. Data on uninoculated alders in hydroponics

Clone Plant Terminal Terminal Terminal stem nutrient leaf ' dry weight solution pH dry weight (g) (g)

1-26 1 6.4 .1812 .3401 1-26 2 6.6 .5284 .4855 2-50 1 6.0 .3741 .4523 2-50 2 6.4 .1962 .3815 3-21 1 6.7 .2184 .6521 3-21 2 6.2 .0781 .6125 1-23 1 6.0 .4244 .3910 1-23 2 6.3 .2009 .3054 5-50 1 6.5 .0804 .4596 5-50 2 6.2 .2385 .7169 3-13 1 6.0 .4024 .5451 3-13 2 6.5 .2790 .4283 4—40 1 6.3 .3138 .5152 4-40 . 2 6.6 .2312 .5061 6-15 1 6.2 .2496 .5289 6-15 2 6.7 .0770 .2910 5-40 1 6.6 .0000 .2669 5-40 2 6.3 .0267 .3799 5-37 1 6.3 .1272 .4012 5-37 2 6.5 .3503 .4367 91

Table A-15. Data on uninoculated alders in hydroponics

Clone Plant Terminal Terminal Terminal root total leaf N dry weight dry weight (g) (g) (g)

1-26 1 .7154 1.2367 .0021 1-26 2 1.0665 2.0804 .0062 2-50 1 1.1175 1.9439 .0051 2-50 2 .9592 1.5369 -0021 3-21 1 .9384 1.8089 .0027 3-21 2 1.3643 2.0549 .0011 1-23 1 1.0187 1.8341 .0061 1-23 2 .8600 1.3663 .0028 5-50 1 1.2079 1.7479 .0009 5-50 2 1.9239 2.8793 .0028 3-13 1 1.0670 2.0145 .0045 3-13 2 .7830 1.4903 .0030 4-40 1 1.3176 2.1466 .0047 4-40 2 .9238 1.6611 .0030 6-15 1 1.4601 2.2386 .0037 6-15 2 .7972 1.1652 .0013 5-40 1 .4201 0.6870 .0000 5-40 2 .5934 1.0000 .0006 5-37 1 .9285 1.4569 .0023 5-37 2 .9783 1.7653 .0063 92

Table A-16. Data on uninoculated alders in hydroponics

Clone Plant Terminal Terminal Terminal stem N root N total N (g) (g) (g)

1-26 1 .0019 .0094 .0134 1-26 2 .0030 .0195 .0287 2-50 1 .0034 .0118 .0203 2-50 2 .0021 .0124 .0166 3-21 1 .0040 .0120 .0187 3-21 2 .0038 .0210 .0259 1-23 1 .0022 .0116 .0199 1-23 2 .0020 .0112 .0160 5-50 1 .0029 .0155 .0193 5-50 2 .0042 .0233 .0303 3-13 1 .0033 .0154 .0232 3-13 2 .0024 .0116 .0170 4-40 1 .0031 .0157 .0235 4-40 2 .0036 .0127 .0193 6-15 1 .0034 .0164 .0235 6-15 2 .0023 .0120 .0156 5-40 1 .0023 .0120 .0156 5-40 2 .0034 .0102 .0142 5-37 1 .0033 .0135 .0191 5-37 2 .0044 .0153 .0260 93

Table A-17. Data on fertilized alders in potting mix

Clone Plant Initial Initial Terminal leaf dry weight total N dry weight estimate (g) estimate (g) (g)

1-26 1 .82 .0191 14.8205 1-26 2 .71 .0168 19.0780 2-50 1 1.53 .0336 25.9121 2-50 2 1.01 .0229 26.4375 3-21 1 .54 .0132 14.9335 3-21 2 1.21 .0271 12.7318 1-23 1 .77 .0180 26.2380 1-23 2 .81 .0188 21.6017 5-50 1 .81 .0188 13.8493 5-50 2 1.07 .0242 11.4562 3-13 1 .81 .0188 15.4260 3-13 2 .79 .0184 8.8795 4—40 1 .74 .0174 11.3753 4—40 2 .98 .0223 16.3590 6-15 1 .75 .0176 6.5667 6-15 2 .45 .0113 6.6463 5-40 1 .29 .0080 1.9968 5-40 2 .66 .0157 5.5373 5-37 1 .42 .0107 3.7283 5-37 2 .67 .0159 2.8977 94

Table A-18. Data on fertilized alders in potting mix

Clone Plant Terminal Terminal Terminal stem root total dry weight dry weight dry weight (g) (g) (g)

1-26 1 9.4833 5.5403 29.8641 1-26 2 17.5723 7.2548 43.9051 2-50 1 23.5297 8.9833 58.4251 2-50 2 24.6763 9.6429 60.7567 3-21 1 11.0338 5.1551 31.1224 3-21 2 10.4918 4.9833 28.2069 1-23 1 24.3846 10.2749 60.8975 1-23 2 17.6906 6.0878 45.3801 5-50 1 9.6671 6.3096 29.8260 5-50 2 6.9113 3.6913 22.0588 3-13 1 11.8361 5.4238 32.6859 3-13 2 7.9377 2.7563 19.5735 4-40 1 8.6404 4.1692 24.1849 4-40 2 13.0674 5.8305 35.2569 6-15 1 4.4831 2.7285 13.7783 6-15 2 5.5992 2.5239 14.7694 5-40 1 .7322 .7376 3.4666 5-40 2 2.1434 1.7204 9.4011 5-37 1 1.6919 1.2210 6.6421 5-37 2 .9835 .8421 4.7233 95

Table A-19. Data on fertilized alders in potting mix

Clone Plant Terminal Terminal Terminal Terminal leaf N stem N root N total N (g) (g) (g) (g)

1-26 1 .6966 .1555 .1524 1.0045 1-26 2 .8624 .2056 .1582 1.2280 2-50 1 1.2360 .3365 .2066 1.7791 2-50 2 1.1183 .3850 .1794 1.6827 3-21 1 .7302 .2085 .1500 1.0887 3-21 2 .4813 .1658 .1435 .7906 1-23 1 1.2306 .3706 .2209 1.8221 1-23 2 .9008 .2901 .1278 1.3187 5-50 1 .6398 .1324 .1237 .8959 5-50 2 .5155 .1120 .0760 .7035 3-13 1 .6911 .1266 .0955 .9132 3-13 2 .3259 .1016 .0562 .4837 4-40 1 .5710 .0873 .0876 .7459 4-40 2 .7787 .1424 .1697 1.0908 6-15 1 .2640 .0623 .0598 .3861 6-15 2 .3024 .0661 .0485 .4170 5-40 1 .0982 .0176 .0221 .1379 5-40 2 .2580 .0459 .0470 .3509 5-37 1 .1883 .0352 .0298 .2533 5-37 2 .1246 .0211 .0206 .1663 96

Table A-20. Data for regression equations to estimate initial dry- weight and nitrogen content of alders

Clone Total Total Total fresh weight dry weight N (g) (g) (g)

1-26 5.4 .82 .0213 1-26 4.2 .71 .0175 2-50 5.3 .85 ' .0194 2-50 9.6 1.53 .0341 3-21 3.0 .54 .0143 1-23 5.1 .82 .0173 1-23 5.7 .77 .0207 5-50 5.3 .81 .0201 5-50 8.1 1.23 .0264 3-13 4.7 .81 .0149 3-13 5.8 .79 .0241 4-40 4.7 .74 .0167 4-40 6.0 .98 .0226 6-15 3.7 .75 .0135 6-15 2.1 .45 .0117 5-40 1.6 .29 .0081 5-37 4.0 .67 .0162 5-37 2.1 .42 .0121 97

Table A-21. Data for regression equation to estimate alder leaf area

Clone Leaf length Leaf width Leaf area (cm) (cm) (cm^)

1-26 3.2 2.7 7.10 1-26 7.4 6.5 35.35 2-50 6. 6 6.5 35.48 2-50 3.1 2.5 6.77 2-50 6.7 6.2 32.25 2-50 5.6 5.5 24.96 2-50 7.0 5.7 30.70 2-50 4.0 4.0 12.90 2-50 1.8 1.5 2.06 3-21 3.4 2.5 6.45 3-21 3.9 3.5 10.97 1-23 3.5 3.2 8.51 1-23 2.9 2.5 6.00 1-23 4.5 4.1 15.48 1-23 4.6 3.6 13.03 5-50 2.6 2.2 4.58 5-50 4.2 4.2 13.93 5-50 1.2 1.0 .77 3-13 4.8 4.0 15.54 3-13 4.4 4.1 14.64 3-13 1.6 1.4 19.35 4-40 2.7 2.2 4.39 4-40 2.1 1.7 2.90 4-40 6.7 5.2 29.73 6-15 4.5 3.9 14.71 6-15 3.5 3.1 9.74 6-15 5.8 4.3 20.64 6-15 4.5 4.9 11.87 5-37 4.6 4.4 17.54 5-37 3.8 3.3 10.97 5-37 3.4 2.5 7.35 5-37 2.8 2.5 6.51 5-37 1.4 1.1 1.16 5-37 1.3 1.2 1.74 5-37 1.7 1.3 2.06