other mineral nutrients. All flasks were inoculated
with the cellulolytic fungus, Chaetomium thermophile
and incubated for 4 weeks at 45°C. At that stage
Chaetomium had degraded 1 g of the original flask
contents but its growth had stopped and there was no
further breakdown after 7 weeks. This was shown to
be due to nitrogen depletion, because the addition of
extra nitrogen at 4 weeks led to further cellulose
breakdown (Fig. 11.13).
Other treatments were also introduced at 4 weeks (the
stage at which Chaetomiumwas nitrogen-depleted)
but no extra nitrogen was supplied. In all cases these
treatments involved placing a small inoculum block
of a thermophilic fungus on the original Chaetomium
colony. When Scytalidium thermophilum was added
to the Chaetomiumflasks at 4 weeks there was no fur-
ther breakdown of the cellulose, although Scytalidium
can degrade cellulose when grown alone. By contrast,
the addition of either Thermoascus aurantiacus or
Aspergillus fumigatusto the 4-week Chaetomiumflasks
led to a further weight loss after 7 weeks. A. fumigatus
was the most effective in this respect – it almost
doubled the original weight loss. These fungi were
seen to grow over the original Chaetomiumcolony. Even
a noncellulolytic fungus, Thermomyces lanuginosus,
caused some additional weight loss when it was added
after 4 weeks.
The implication of these findings is that the early
colonizers in a substrate succession (in this case
Chaetomium which has the highest temperature
optimum for growth) can be displaced by fungi that
occur later in the succession (e.g. Thermoascusor A.
fumigatus) in conditions of nitrogen starvation.
Basidiomycota (Agaricus bisporus, Coprinus cinereus, etc.)
typically occur late in the succession of fungal activit-
ies in natural materials, and many of them have been
shown to use proteins as nitrogen sources. Indeed, A.
bisporuscan grow in culture media when nitrogen is
supplied only in the form of living or heat-killed bac-
teria. We noted earlier that Phanerochaeteis induced
to synthesize lignin-degrading enzymes in response to
nitrogen limitation; and in Chapter 12 we will see that
Basidiomycota such as Coprinusdisrupt the hyphae
of other fungi on contact, which perhaps provides a
source of organic nitrogen. So it seems that nitrogen
availability is a key factor in fungal successions, and
that the later colonizers have special abilities to obtain
(recycle) the nitrogen that earlier colonizers have
utilized.
Fungal decomposers in the root zone
Living roots provide a continuous input of nutrients
into soil, evidenced by the fact that motile pseudomo-
nads and zoospores of Oomycota or plasmodiophorids
accumulate near the root tips. Amino acids, sugars,
and other organic molecules are found frequently in
root exudates, and the total microbial population
increases progressively with distance behind the tips
until it reaches a plateau. At this point the population
level is likely to represent the “carrying capacity” of the
root – in other words, the rate of continuing nutrient
release is matched by the rate at which these nutrients
are utilized by the existing population.
The zone of soil influenced by the presence of a root
is termed the rhizosphere. As we saw in Chapter 10,
fungal spores can be induced to germinate by the
presence of root exudates or even by small signalling
molecules that may not serve as nutrients. But this
is only the start of a continuous process of root
colonization by saprotrophs, parasites, and pathogens
until, eventually, the roots die and their nutrients are
returned to the soil.
The behavior of roots is difficult to investigate in
field conditions, but this can be done in a glasshouse
if plants are grown in soil containers with a sloping
transparent face so that periodic observations can
be made (the roots being blacked out except during
periods of observation). Figures 11.14 and 11.15 show
the results of one such experiment where groundnut
plants (Arachis hypogea) were grown over a 14-week
period (Krauss & Deacon 1994). The whole visible part
of the root system was traced at weekly intervals, and
the individual roots were scored as being either alive
(white roots) or dead (brown decaying roots or roots
that had disappeared).
Figures 11.14 and 11.15 reveal an astonishingly
high rate or rhizodeposition, i.e. the shedding of root
material into the soil where it becomes available to
the microbial population of the rhizosphere. Although
the total (cumulative) root length increased through-
out the period of observation, up to plant maturity at
14 weeks or 20 weeks (depending on the groundnut
cultivar), the maximum length of living (white)
rootson any plant was reached at between 2 and
4 weeks after sowing the seeds. Beyond that time, the
taproot continued to grow and produced more root
laterals, but the rate of root death (disappearance)
exceeded the rate of new root production, leading to
a decline in total root length. An early onset of root
lateral death, 5 weeks after sowing, was also recorded
in a field plot of groundnuts in Malawi. So, at least for
this crop, there is a continuous process of “root shed-
ding” which would provide a continuous input of
organic matter for fungi and other soil microbes.
Other studies have focused on the roots of cereals
and grasses, using vital dyes to follow the progressive
death of the root cortex. Acridine orangepenetrates
roots, binds to the DNA in the nuclei, and fluoresces
bright green under a fluorescence microscope (Fig.
11.16a,c). Alternatively, roots can be infiltrated with