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Appl Environ Microbiol, April 1998, p. 1430-1435, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Detection of Infectious Tobamoviruses in
Forest Soils
Ronald C.
Fillhart,
George D.
Bachand, and
John D.
Castello*
Faculty of Environmental and Forest Biology,
College of Environmental Science and Forestry, State University of
New York, Syracuse, New York 13210-2788
Received 3 July 1997/Accepted 24 September 1997
 |
ABSTRACT |
Our objectives were to evaluate elution and bait plant methods to
detect infectious tobamoviruses in forest soils in New York State.
Soils were collected from two forest sites: Whiteface Mountain (WF) and
Heiberg Forest (HF). The effectiveness of four buffers to elute tomato
mosaic tobamovirus (ToMV) from organic and mineral fractions of WF soil
amended with ToMV was tested, and virus content was assessed by
enzyme-linked immunosorbent assay (ELISA). The effectiveness of
Chenopodium quinoa (Willd.) bait plants to detect the virus
also was tested. Both methods then were utilized to detect
tobamoviruses in 11 WF and 2 HF soil samples. A phosphate buffer (100 mM, pH 7.0) eluted more ToMV from soil than the other buffers tested.
Mineral soil bound more virus than organic soil. Virus recoveries from
virus-amended organic and mineral soils were 3 and 10%, respectively,
and the detection sensitivity was 10 to 20 ng/g of soil. Roots of bait
plants grown in all virus-amended soils tested positive by ELISA, and
virus concentrations averaged 10 ng/g. Both ToMV and tobacco mosaic
tobamovirus (TMV) were transmitted to C. quinoa by elution
from one of two HF soil samples but not from the WF soil samples. A
tobamovirus was detected by bait planting in 12 of 73 (16%) root
extracts representing 5 of 13 soil samples (38%). Tobamovirus-like
particles were seen by transmission electron microscopy in 6 of 12 infected root extracts. Tobamoviruses occur in forest soils in New York
State. Abiotic soil transmission to trees may permit localized spread
and persistence of these viruses in forest ecosystems.
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INTRODUCTION |
Plant viruses occur in natural
ecosystems (13) including the air (10), water
(8), soil (7), and trees (28) of forests. Many of the viruses detected in forest ecosystems lack biotic
vectors and are soilborne (abiotic soil transmission), e.g., tobamo-,
potex-, and tombusviruses. However, the ecological and epidemiological
significance of abiotic, soilborne virus transmission is unknown
(23). Soilborne transmission of tomato mosaic tobamovirus (ToMV) and tobacco mosaic tobamovirus (TMV) in agricultural or greenhouse settings has been investigated (2, 5, 6, 12, 26,
31), but studies in natural ecosystems are lacking. Although such
studies may help to explain how these and similar viruses spread into
and/or within agricultural and forest ecosystems, very little research
has been conducted on the development of sensitive methods for
detecting infectious plant viruses in soils.
ToMV is one of 13 plant viruses that spread under natural conditions
without a biotic vector (17, 23). ToMV is soilborne (17) and waterborne (19), has a wide host range
(17), is extremely stable, and infects plants through roots
(30, 31). Recently, it has been detected in red spruce
(Picea rubens Sarg.) (21), stream water
(19), and clouds (10) in high-elevation, montane,
spruce-fir ecosystems. The virus infects red spruce by an airborne
mechanism, and infected trees occur throughout the northeastern United
States (15). Red spruce seedlings become infected when
inoculated with purified virus (4, 20). However, the virus
has not been detected in soil in which infected red spruce trees are
found, and the specific infection mechanism is unknown.
ToMV infection affects both the growth and physiology of red spruce.
Infection of dominant and codominant red spruce trees on Whiteface
Mountain, N.Y. (WF), was positively correlated with the number of fine
roots and negatively correlated with the length of the live crown
(11). Infection of seedlings caused a 50% reduction in rate
of increase of height, weight, and root volume compared with that of
noninfected seedlings, but the freezing tolerance of the infected
seedlings was greater than that of noninfected seedlings
(4).
To our knowledge, only one study has attempted to detect soilborne
plant viruses in forest ecosystems. Büttner and Nienhaus (7) transmitted plant viruses in the potex-, tobamo-,
necro-, and potyvirus genera from German forest soils to herbaceous
and/or woody hosts. About one-third of 284 soil samples tested were
positive for one or more of the viruses. Because many European forests occur on what was once agricultural land, the viruses detected in these
trees and soils may have originated from infected crop residues. As
more forests develop on former agricultural land (3, 29),
the incidence of virus diseases in forest tree species in the United
States may increase. Alternatively, viruses detected in forest
ecosystems with little or no prior history of agriculture may be
indigenous or introduced by some other mechanism. In either case,
knowing what viruses are present in forest ecosystems, how best to
detect them, their origin, how they spread, and their effects on tree
growth and development will be necessary to assess their impact and to
develop effective disease management strategies. Thus, the objectives
of this study were to evaluate methods to detect infectious
tobamoviruses and to demonstrate their presence in forest soils. We
hypothesize that infectious tobamoviruses can be detected by elution
and bait plant methods in forest soils collected from stands where
virus-infected trees occur.
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MATERIALS AND METHODS |
Soil collection.
The research was conducted primarily with
soil collected from WF (northwestern slope at an elevation of 1,070 m,
44°22'55"N and 73°54'30"W), because our interest lies in the
epidemiology and ecology of this virus in high-elevation, montane,
spruce-fir ecosystems. However, for comparative purposes, two soil
samples also were collected from Heiberg Memorial Forest, Cortland
County, N.Y. (HF) (42°46'45"N and 76°5'11"W), and assayed for ToMV.
ToMV-infected red spruces also are present on the HF site
(8a), and the virus has been detected in stream water there
(19). In addition, TMV-infected white ash (Fraxinus
americana L.) can be found at this site (9). The two
sites differ in disturbance history. WF was logged from the late 1800s
to the early decades of this century and experienced slash fires and
soil erosion until the 1930s. HF was logged earlier and was used for
agriculture and pasture until the 1950s, at which time the site
reverted to northern hardwood forest (e.g., American beech, yellow
birch, and sugar maple). The two sites also differ in soil type
(14). WF soil is a Spodosol of the Sisk-Glebe complex consisting of silt loam overlaid by an organic horizon. HF soil is a
Mardin channery silt loam. Soil pH, cation-exchange capacity, texture,
percent organic matter for HF soil, the organic and mineral fractions
for WF soil, and Promix potting mixture (Premier Horticulture, Inc.,
Rivière-du-Loup, Quebec, Canada) were as reported by Fillhart (14).
Soil from WF was collected from three soil pits in June 1994 and
February, April, August, September, and December 1995. WF soil was
separated into mineral and organic fractions. Soil was collected from
beneath infected red spruce at HF in February and September 1995, upslope from a small stream in which ToMV also was previously isolated
(19). All soils were stored moist at 4°C because soil
drying can inactivate tobamoviruses (2, 12, 36).
Elution methodology.
Two methods to detect ToMV in soil were
evaluated: elution and the use of bait plants. Both methods
subsequently were utilized to detect the virus in forest soils.
For the elution methodology, the effectiveness of four buffers to elute
a WF stream water isolate of ToMV (ToMV-38) from the
mineral and
organic fractions of WF soil was tested: 100 mM citrate
buffer (pH
3.0), 100 mM citrate buffer (pH 5.0), 100 mM phosphate
buffer (pH 7.0),
and 100 mM borate buffer (pH 9.0). These buffers
were selected to
provide a range of pH values because virus elution
is highly dependent
on the pH of the elution buffer used (
18).
Two micrograms of
purified ToMV in 1 ml of autoclaved, deionized
water was added to
organic (3 g) and mineral (5 g) soil samples
in separate 125-ml flasks.
The flasks were shaken by hand for
5 min. Approximately 25 ml of
autoclaved, deionized water was
added to each flask, shaken at 250 rpm
for 1 h, and incubated
overnight at 4°C. The solutions were
filtered under a vacuum through
Whatman no. 1 filter paper, and the
filtrates were centrifuged
for 2 h at 100,000 ×
g. Pellets were suspended in 500 µl of double-antibody
sandwich, enzyme-linked immunosorbent assay (DAS-ELISA) extraction
buffer (EEB) (
20). A 25-ml aliquot of test buffer then was
added
to the soil in each flask, shaken, incubated, filtered, and
centrifuged
as described above. Pellet suspensions obtained from the
water
and buffer eluates were tested separately for ToMV by DAS-ELISA
as described by Jacobi and Castello (
20). Organic and
mineral
fractions without added virus were treated similarly to serve
as negative controls for DAS-ELISA.
The protocol described above is comprised of elution and
ultracentrifugation phases. The percentage of virus recovered from
each
phase of the protocol was determined as follows. For the
elution phase,
1 g of air-dried mineral soil or 100 mg of organic
soil was placed
into separate 1.5-ml microcentrifuge tubes. Purified
ToMV was added in
amounts of 0, 1, 5, 10, 25, 50, 100, 250, 500,
1,000, or 1,500 ng in 1 ml of 100 mM phosphate buffer (pH 7.0).
The suspensions were agitated
on a vortex mixer for 1 min, shaken
on a rotary shaker (250 rpm) for 30 min, incubated at 4°C overnight,
and then centrifuged at 10,000 ×
g for 2 min. The supernatants
were tested for ToMV by
DAS-ELISA. The amount of added virus that
was recovered was calculated
by comparing the absorbance at 405
nm (
A405)
with a dilution series of purified virus. The percent
virus recovery
was calculated as follows: (nanograms of virus
recovered/nanograms of
virus added) × 100.
For the ultracentrifugation phase, 1 ml of purified ToMV was added to
separate flasks each containing 10 ml of 100 mM phosphate
buffer (pH
7.0) for final concentrations of 1, 5, 10, 15, 20,
25, 50, and 100 ng
of ToMV in 11 ml. The solutions were filtered
under vacuum through
Whatman no. 1 filter paper and washed with
15 ml of buffer for a total
of 26 ml/flask. The virus-containing
filtrates were centrifuged at
100,000 ×
g for 2 h. The resulting
pellets were
suspended in 1 ml of EEB and tested for ToMV by DAS-ELISA.
The amount
and percentage of virus recovered were calculated as
described above.
The percent virus recovery in both phases combined
was determined as
follows: (percent recovery in the elution phase
× percent
recovery in the centrifugation phase)/100.
The complete elution protocol was tested as follows: 5-g aliquots of WF
mineral soil and 3-g aliquots of WF organic soil were
placed into
separate 125-ml flasks. A 2-ml aliquot of autoclaved,
deionized water
containing purified ToMV in amounts of 0, 50,
100, 250, 500, 1,000, or
1,500 ng was added to separate flasks
and subjected to the complete
elution protocol described above.
The resultant water and buffer
eluates were tested for ToMV by
DAS-ELISA, and the percent virus
recovery was determined as described
above.
To determine the effect of soil extracts on the infectivity of ToMV,
dilutions of purified virus were prepared in WF mineral
and organic
soil fractions as follows. One liter of 100 mM phosphate
buffer (pH
7.0) was added to 1 kg of mineral soil and to 500 g
of organic
soil, and the slurries were shaken at 250 rpm for 1
h followed by
incubation overnight at 4°C. The slurries were filtered
through
Whatman no. 1 filter paper under a vacuum, and the filtrates
were
centrifuged at 100,000 ×
g for 2 h. The pellets
were then
suspended in 1 ml of 10 mM phosphate buffer (pH 7.0).
Dilutions
of purified ToMV (0, 1, 10, 100, and 1,000 ng/ml) were
prepared
in the pellet resuspensions from mineral and organic soils and
in 10 mM phosphate buffer (pH 7.0). Celite was added to each of
the
preparations, which were then used to inoculate leaves of
Chenopodium quinoa Willd. Lesions were counted after a 5-day
incubation
in the greenhouse.
Bait plant methodology.
To determine whether C. quinoa plants would become infected if grown in soil containing
infectious virus, the basic method of Büttner and Nienhaus
(7) was used, modified as described here. Seeds were planted
in three 15-cm-diameter pots containing WF organic soil, WF mineral
soil, or Promix. Ten grams of fresh, symptomatic tobacco
(Nicotiana tabacum L. cv. Turkish) tissue systemically
infected with ToMV was chopped, added to each pot, and mixed
thoroughly. As a control, seeds were planted in pots containing soils
amended with noninfected tobacco tissue. Upon germination, plants were
thinned to 15 per pot and grown in the greenhouse with a day length of
15 h for 6 weeks. Roots were tested for ToMV by DAS-ELISA.
Detection of tobamoviruses in forest soils.
The elution and
bait plant techniques then were used to detect infectious tobamoviruses
in forest soils. For the elution protocol, 1 liter of 100 mM phosphate
buffer (pH 7.0) was added to 1 kg each of WF mineral and HF soils or to
500 g of WF organic soil obtained at each collection date. The
resultant slurries were treated as described above. The pellets from
high-speed centrifugation were suspended in a total volume of 1 ml of
10 mM phosphate buffer (pH 7.0) per soil sample. C. quinoa
plants then were mechanically inoculated with each suspension and
monitored for symptom development for 2 weeks. Thirteen soil samples
(11 and 2 samples from WF and HF, respectively) were assayed for
infectious virus in this manner.
The same 13 soil samples also were tested for infectious ToMV by the
bait plant protocol modified as follows: soils were mixed
1:1 (vol/vol)
with autoclaved Promix because plant growth was
poor in unamended
soils. To duplicate soil and water conditions
on WF, plants were
watered with water collected from Esther Brook
on WF and autoclaved.
The plants were thinned to 20 per pot at
1 week after germination and
were harvested 8 weeks after planting.
Roots of two to six plants per
pot were collected and assayed
separately for ToMV by DAS-ELISA. In
addition, roots from the
remaining 14 to 18 plants in each pot were
combined, triturated
in 10 mM phosphate buffer (pH 7.2), clarified by
low-speed centrifugation
(10,000 ×
g for 10 min),
concentrated by ultracentrifugation (100,000
×
g for
2 h), suspended in 500 µl of 10 mM phosphate buffer (pH
7.2),
and used to inoculate the leaves of
C. quinoa, which were
then monitored for symptoms for 2 weeks. Approximately 250 g of
soil from each pot then was leached with 600 ml of 100 mM phosphate
buffer (pH 7.0) as described above, and the suspended pellets
were used
to inoculate the leaves of
C. quinoa.
TEM.
Transmission electron microscopy (TEM) was conducted on
leaf extracts of C. quinoa plants that showed symptoms when
inoculated with concentrated soil eluates and in root extracts of bait
plants in which the virus was detected by DAS-ELISA. Formvar and
carbon-coated grids for TEM were incubated for 15 min on crude tissue
extracts, rinsed with 100 mM phosphate buffer (pH 7.0) and distilled
water, negatively stained with 2% aqueous uranyl acetate, air dried, and viewed on a Japanese Electron Optics Laboratory JEM 2000EX transmission electron microscope.
| |
RESULTS |
Elution methodology.
More ToMV bound to WF mineral than to
organic soil because water alone removed more than 100 ng of ToMV from
virus-amended organic soil and approximately 50 ng from virus-amended
mineral soil (Table 1). The 100 mM
phosphate buffer (pH 7.0) eluted more ToMV from both mineral and
organic soils than did the other buffers tested (Table 1). By
comparison to the A405 values of a dilution series with purified virus, more than 100 ng and approximately 75 ng of
ToMV were eluted from mineral and organic soils, respectively (Table
1). Increasing the molarity (0.2 M) or adding Tween 80 (0.1%) to
phosphate buffer did not increase the amount of virus eluted from these
soils (data not shown).
The percent recovery of ToMV from the elution phase ranged from 8 to
30%, with a mean of 17% for organic and 22% for mineral
soils (Table
2). The percent recovery of ToMV from the
centrifugation
phase ranged from approximately 20 to 30%, with a mean
of 26%
(Table
3). When both phases of
the protocol are combined, approximately
5% of the ToMV added to WF
soils should be recoverable, which
agrees well with the calculated
recoveries of ToMV from virus-amended
organic and mineral soils of 3 and 10%, respectively (Table
4).
Based
upon the ELISA results, the detection sensitivity of the
elution method
was approximately 10 to 20 ng of ToMV/g of soil
(Table
4).
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TABLE 2.
Amounts and percentages of ToMV recovered from
unconcentrated eluates of virus-amended mineral and organic WF soils
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Dilution of purified ToMV in mineral and organic soil extracts reduced
virus infectivity.
C. quinoa plants inoculated with
1 ng of
ToMV per ml prepared in phosphate buffer produced 23 local
lesions,
whereas no lesions were produced on plants inoculated
with virus
prepared in mineral or organic soil extracts. Plants
inoculated with
10-ng/ml samples of virus prepared in buffer,
organic soil, and mineral
soil extracts produced 200, 30, and
3 lesions, respectively, a finding
which corresponds to 85 and
98% inhibition of infectivity,
respectively. All plants inoculated
with 100- and 1,000-ng/ml samples
of virus produced numerous local
lesions.
Bait plant methodology.
Plant survival and growth in the WF
soils were poor, particularly in the mineral soil. Therefore, the roots
of surviving plants grown in mineral and organic soils were combined by
soil type to obtain sufficient tissue for DAS-ELISA. Roots collected
from plants grown in all virus-amended soils tested positive by
DAS-ELISA (the mean A405 values of infected and
control root samples were 1.00 and 0.14, respectively). The composite
root sample from plants grown in non-virus-amended organic soil, but
not Promix or mineral soil, also tested positive for ToMV by DAS-ELISA
(the A405 values of the infected root sample and
the control root sample were 0.27 and 0.14, respectively). When the
absorbances were compared with those of dilutions of purified virus,
the mean virus concentration within infected roots was found to be
approximately 10 ng/g.
Tobamovirus detection in forest soils.
Eleven necrotic local
lesions developed on C. quinoa leaves inoculated with the
concentrated eluate from HF soil collected in February 1995 (Table
5). Many rigid rods approximately 250 to
300 nm in length and 18 nm in diameter were observed by TEM of infected
leaf tissue. Black turtle beans (Phaseolus vulgaris L. cv.
Black Turtle 1), a differential host for TMV and ToMV, were
mechanically inoculated separately with several of the lesions that
developed on this C. quinoa plant. Some bean plants
developed red, necrotic, local lesions characteristic of TMV, and
others did not produce symptoms, which indicated the presence of ToMV but not TMV. Symptoms were not observed on C. quinoa plants
inoculated with other concentrated soil eluates.
A tobamovirus was detected by DAS-ELISA in the roots of
C. quinoa plants grown in WF and HF soils (Table
5). There was no
relationship between virus detection and the time of soil collection.
Virus was detected in soils collected in June and August (WF organic),
December (WF mineral), and February and September (HF) (Table
5).
Several tobamovirus-like particles (250 to 300 nm by 18 nm)
were
detected by TEM in 6 of the 12 ELISA-positive
C. quinoa root
extracts. Tobamoviruses were not transmitted to
C. quinoa
from
root or soil concentrates.
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DISCUSSION |
A tobamovirus was detected in HF and WF soils by DAS-ELISA of bait
plant roots and then confirmed by TEM in 50% of those samples (Table
5). Because of the cross-reactivity of ToMV antiserum, the ToMV
DAS-ELISA cannot distinguish between TMV and ToMV in infected tissues.
Therefore, the identity of the virus detected in soil cannot be
determined. However, because ToMV has been detected in clouds
(10), red spruce (21), and stream water
(19) on WF, we believe that the virus detected in WF soil is
ToMV and not TMV. However, based on particle morphology and the
presence or absence of symptoms in the differential host bean cultivar Black Turtle 1, both TMV and ToMV were transmitted to C. quinoa from one concentrated HF soil eluate (Table 5). Both ToMV
and TMV have been detected previously at the HF site, with ToMV being detected in stream and pond water (19) and in red spruce
(8a) and TMV being detected in white ash (F. americana L.) (9). These results support our hypothesis
that infectious tobamoviruses are present in soils of forest stands in
which infected trees occur and, to our knowledge, represent only the
second report of infectious plant viruses detected in forest soils.
Although both the elution and bait plant methods permitted detection of
infectious tobamoviruses in forest soils, neither TMV nor ToMV was
detected frequently. Both viruses were transmitted from only 1 of 13 concentrated soil eluates (Table 5), and they were detected by ELISA in
only 12 of 73 (16%) bait plant root extracts representing 5 of 13 soil
samples (38%) (Table 5). Apparently, the bait plant method is more
sensitive than elution for the detection of tobamoviruses in forest
soils. Although plant viruses in six different genera have been
detected in soil using the bait plant technique (16),
tobamoviruses were rarely (5%) detected by this method in forest soils
in Germany (7). Tobamoviruses may not be common in forest
soils or their concentration may be too low to permit reliable
detection by elution-infectivity bioassay. Similarly, exposure of
C. quinoa roots to low levels of virus inoculum in forest
soil may be too short to generate more than a few localized root
infections. Long-term exposure to a low-concentration inoculum may be
required to initiate infection or to achieve a high virus concentration
within infected tissues. However, because of its extremely long life
span (400 years), red spruce may be more likely to become infected when
growing in soil containing low levels of virus. Therefore, soilborne
virus may be a source of inoculum for infection of this and other
long-lived woody species. The duration and degree of host exposure to
inoculum, as well as the inoculum concentration, directly influence
abiotic, soilborne virus transmission (16). Likewise, the
incidence of ToMV infection of tomato plants decreased with decreased
inoculum concentration (31). Methods more sensitive than
elution or bait plants followed by infectivity bioassay or ELISA are
now available for virus detection. DAS-ELISA and PCR amplification can
detect nanogram and femtogram quantities of virus, respectively, but
these methods are problematic for direct use with soils that often
contain interfering substances (1). We have recently
developed a multiplex immunocapture reverse transcriptase PCR protocol
(22), which should permit detection and differentiation of
TMV and ToMV with great sensitivity directly in the roots of bait
plants or clarified soil eluates.
Techniques to extract, concentrate, and enumerate infectious viruses
from soils come primarily from elution studies with human and animal
viruses. These techniques are marginally efficient for a few
enteroviruses and inadequate for others (34). Elution followed by direct assay on tissue-cultured animal cell lines is used
routinely to detect animal and human viruses in soils (18,
39). For example, recovery of poliovirus from estuarine sediments
ranged from 0.1 to 71% (mean = 18% [39]), and
the recovery of four enteroviruses from a loamy sand soil ranged from 0.1 to 88% (18). By comparison, the recoveries of ToMV by
elution from organic and mineral soils were 19 and 22%, respectively
(Table 2). Unfortunately, only 25% of the eluted virus was recovered during the subsequent centrifugation phase (Table 3). In the only study
of which we are aware that utilized elution to detect plant viruses in
soil, Cheo (12) eluted TMV from agricultural soil with 25 mM
phosphate buffer (pH 7.5) followed by differential centrifugation and
either infectivity bioassay or spectrophotometry to detect the virus.
Detection sensitivity was low: 10 to 1,000 ng/ml for the infectivity
bioassay and 50 to 300 µg/ml for spectrophotometry. The sensitivity
of our infectivity bioassay to detect ToMV was approximately 10 ng/ml
for virus prepared in soil extracts, which was 10-fold less than the
sensitivity of the assay to detect virus prepared in phosphate buffer.
Humic acids in organic soils are known to inhibit the infectivity of
TMV in tobacco (33). Therefore, assuming a 10% recovery of
virus by elution (Table 4), at least 100 ng of virus must be present in
a soil sample in order for it to be detected by elution and infectivity
bioassay.
Tobamoviruses persist in plant debris in soil (26).
Therefore, their presence in forest soils and trees may relate to past land use history. Many European forests, in which viruses have been
detected (7), were planted on former agricultural land. The
HF site in this study was in agriculture and pasture prior to 1955. Interestingly, on WF ToMV but not TMV has been detected in trees
(21), stream water (19), clouds (10),
and possibly soil (Table 5). The apparent absence of TMV at this site
may relate to the absence of a prior history of agriculture.
Alternatively, the concentration and persistence of certain viruses in
forest soils may relate to differences in soil or virus characteristics that affect mobility and persistence.
Many factors affect the mobility and persistence of viruses in soils,
including soil pH and moisture content, concentration of ions,
cation-exchange capacity, type and amount of clay, organic matter
concentration, proteins, salt concentrations in soil and groundwater,
hydraulic conditions, infiltration rate, virus type and strain, and
isoelectric point (IEP) (24, 27, 37, 38). More ToMV adsorbed
to WF mineral than to organic soils (Tables 1 and 4). Organic soils
tend not to bind as much virus as mineral soils (34).
Therefore, if virus input is high, ToMV is more likely to remain
unbound in organic soils than in the mineral horizon where clay
particles may bind the virus. Adsorption of some plant (23, 27,
32) and animal (27) viruses to clay particles may
stabilize infectivity. However, adsorption of TMV to bentonite clay
increased its degradation (27). The IEP of the WF stream
isolate is 3.7 (14), which is lower than that of other ToMV
strains (17) and equivalent to that of some TMV strains. pH
determines both type and strength of virus surface charge
(34), such that virions with no net or a weak surface charge
are readily mobilized in soil (38). The IEP of the WF isolate of ToMV is equivalent to the pH of WF organic soil (3.7) and
lower than that of WF mineral soil (4.4) and HF soil (4.8) (14). At soil pH values above the IEP of a virus, as in WF
mineral soil and HF soil, adsorption to clay particles may be
considerably reduced due to the repulsive forces of negatively charged
particles (38). In conjunction with low soil substratum
permeability and rapid surface runoff on the WF site, ToMV may readily
leach from WF soils. In addition, because the IEP of the virus is
identical to that of WF organic soil, slight changes in groundwater or
rainwater pH may alter its mobility in organic soil. Organic soils
contain little clay to bind virus, and humic and fulvic acids interfere with potential binding sites (37). These factors may help to explain why tobamoviruses were not readily detected by direct elution
of WF soils.
Tobamoviruses are present in forest soils in New York State, from which
they may infect forest trees. Both TMV and ToMV have been detected in
forest, shade, and ornamental tree species (9, 11, 25, 28,
35), which could then serve as reservoir hosts to maintain these
viruses in forest ecosystems. Subsequent, abiotic soil transmission to
trees may provide the mechanism for the localized spread of these
viruses and their persistence in forest ecosystems.
| |
ACKNOWLEDGMENTS |
This work was supported by grant 93-37101-8837 to J. D. Castello from the USDA-NRICGP.
We thank D. Wolfe of the Atmospheric Sciences Research Center at WF for
help in the location of soil collection sites and J. Hudyncia for help
with soil collection and elution.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: State University
of New York, College of Environmental Science & Forestry, Faculty of
Environmental & Forest Biology, 1 Forestry Dr., Syracuse, NY 13210. Phone: (315) 470-6789. Fax: (315) 470-6934. E-mail:
jdcastel{at}syr.edu.
| |
REFERENCES |
| 1.
|
Abbaszadegan, M.,
M. S. Huber,
C. P. Gerba, and I. L. Pepper.
1993.
Detection of enteroviruses in groundwater with the polymerase chain reaction.
Appl. Environ. Microbiol.
59:1318-1324[Abstract/Free Full Text].
|
| 2.
|
Allen, W. R.
1984.
Mode of inactivation of TMV in soil under dehydrating conditions.
Can. J. Plant Pathol.
6:9-16.
|
| 3.
|
Anonymous.
1995.
.
The empire state's forests trends in a robust resource. Northeast Forest Experiment Station report NE-INF-126-95.
USDA Forest Service, Washington, D.C.
|
| 4.
|
Bachand, G. D.,
J. D. Castello,
M. Schaedle,
S. V. Stehman, and W. H. Livingston.
1996.
Effects of tomato mosaic tobamovirus infection on red spruce seedlings.
Can. J. For. Res.
26:973-981.
|
| 5.
|
Broadbent, L.
1965.
The epidemiology of tomato mosaic. VII. Virus infection through tomato roots.
Ann. Appl. Biol.
55:57-66.
|
| 6.
|
Broadbent, L.,
W. H. Read, and F. T. Last.
1965.
The epidemiology of tomato mosaic. X. Persistence of TMV-infected debris in soil, and the effects of soil partial sterilization.
Ann. Appl. Biol.
55:471-483.
|
| 7.
|
Büttner, C., and F. Nienhaus.
1989.
Virus contamination of soils in forest ecosystems of the Federal Republic of Germany.
Eur. J. For. Pathol.
19:47-53.
|
| 8.
|
Büttner, C.,
V. Jacobi, and R. Koenig.
1987.
Isolation of carnation Italian ringspot virus from a creek in a forested area southwest of Bonn.
J. Phytopathol.
118:131-134.
|
| 8a.
| Castello, J. D. Unpublished data.
|
| 9.
|
Castello, J. D.,
L. A. Amico, and M. T. O'Shea.
1984.
Detection of tobacco mosaic and tobacco ringspot viruses in white ash trees by enzyme-linked immunosorbent assay.
Plant Dis.
68:787-790.
|
| 10.
|
Castello, J. D.,
D. K. Lakshman,
S. M. Tavantzis,
S. O. Rogers,
G. D. Bachand,
R. J. Jagels,
J. Carlisle, and Y. Liu.
1995.
Detection of infectious tomato mosaic tobamovirus in fog and clouds.
Phytopathology
85:1409-1412.
|
| 11.
|
Castello, J. D.,
P. M. Wargo,
V. Jacobi,
G. D. Bachand,
D. R. Tobi, and M. A. M. Rogers.
1995.
Tomato mosaic virus infection of red spruce on Whiteface Mountain, New York: prevalence and potential impact.
Can. J. For. Res.
25:1340-1345.
|
| 12.
|
Cheo, P. C.
1980.
Antiviral factors in soil.
Soil Sci. Soc. Am. J.
44:62-67.
[Abstract/Free Full Text] |
| 13.
|
Cooper, J. I., and F. O. MacCallum.
1984.
.
Viruses and the environment.
Chapman and Hall, London, England.
|
| 14.
|
Fillhart, R. C.
1996.
.
Studies of the modes of transmission and spread of tomato mosaic tobamovirus in a spruce/fir ecosystem. M.S. thesis.
State University of New York, Syracuse.
|
| 15.
|
Fillhart, R. C.,
G. D. Bachand, and J. D. Castello.
1997.
Airborne transmission of tomato mosaic tobamovirus and its occurrence in red spruce in the northeastern United States.
Can. J. For. Res.
27:1176-1181.
|
| 16.
|
Hiruki, C., and D. S. Teakle.
1987.
Soil-borne viruses of plants.
Curr. Top. Vector Res.
3:177-215.
|
| 17.
|
Hollings, M., and H. Huttinga.
1976.
.
Tomato mosaic virus. Descriptions of plant viruses, no. 156.
Commonwealth Mycological Institute/Association for Applied Biology, Kew, England.
|
| 18.
|
Hurst, C. J., and C. P. Gerba.
1979.
Development of a quantitative method for the detection of enteroviruses in soil.
Appl. Environ. Microbiol.
37:626-632[Abstract/Free Full Text].
|
| 19.
|
Jacobi, V., and J. D. Castello.
1991.
Isolation of tomato mosaic virus from waters draining forest stands in New York State.
Phytopathology
81:1112-1117.
|
| 20.
|
Jacobi, V., and J. D. Castello.
1992.
Infection of red spruce, black spruce, and balsam fir seedlings with tomato mosaic virus.
Can. J. For. Res.
22:919-924.
|
| 21.
|
Jacobi, V.,
J. D. Castello, and M. Flachmann.
1992.
Isolation of tomato mosaic virus from red spruce.
Plant Dis.
76:518-522.
|
| 22.
|
Jacobi, V.,
G. D. Bachand,
R. C. Hamelin, and J. D. Castello.
1997.
Development of immunocapture PCR assays for tobacco and tomato mosaic tobamoviruses.
Phytopathology
87:S46. (Abstract.)
|
| 23.
|
Jones, A. T.
1993.
Virus transmission through soil and by soil-inhabiting organisms.. In
R. E. F. Matthews (ed.), Diagnosis of plant virus diseases.
CRC Press, Inc., Boca Raton, Fla.
|
| 24.
|
Kegler, H.,
E. Fuchs,
D. Spaar, and J. Kegler.
1995.
Viren im Boden und Grundwasser (Übersichtsbeitrag).
Arch. Phytopathol. Pflanzenschutz
29:349-371.
|
| 25.
|
Lana, A. F.,
O. T. Thomas, and J. F. Peterson.
1980.
A virus isolated from sugar maple.
Phytopathol. Z.
97:214-218.
|
| 26.
|
Lanter, J. M.,
J. M. McGuire, and M. J. Goode.
1982.
Persistence of tomato mosaic virus in tomato debris and soil under field conditions.
Plant Dis.
66:552-555.
|
| 27.
|
Lipson, S. M., and G. Stotzky.
1987.
Interactions between viruses and clay minerals, p. 197-230. In
V. C. Rao, and J. L. Melnick (ed.), Human viruses in sediments, sludges and soils.
CRC Press, Inc., Boca Raton, Fla.
|
| 28.
|
Nienhaus, F., and J. D. Castello.
1989.
Viruses of forest trees.
Annu. Rev. Phytopathol.
27:165-186.
|
| 29.
|
Nyland, R. D.,
W. C. Zipperer, and D. B. Hill.
1986.
The development of forest islands in exurban central New York State.
Landsc. Urban Plann.
13:111-123.
|
| 30.
|
Pares, R. D.,
L. V. Gunn, and G. C. Cresswell.
1992.
Tomato mosaic virus infection in a recirculating nutrient solution.
J. Phytopathol.
135:192-198.
|
| 31.
|
Pares, R. D.,
L. V. Gunn, and E. N. Keskula.
1996.
The role of infective plant debris, and its concentration in soil, in the ecology of tomato mosaic tobamovirus, a non-vectored plant virus.
J. Phytopathol.
144:147-150.
|
| 32.
|
Piazzolla, P.,
F. Palmieri, and M. Nuzzaci.
1989.
Infectivity studies on cucumber mosaic virus treated with a clay material.
J. Phytopathol.
127:291-295.
|
| 33.
|
Polak, Z., and F. Pospisil.
1995.
Alleviation of plant virus infection by humic acids.
Biol. Plant.
2:315-317.
|
| 34.
|
Rao, V. C., and J. L. Melnick.
1986.
.
Environmental virology.
American Society for Microbiology, Washington, D.C.
|
| 35.
|
Reddick, B. B.
1989.
Isolation and partial characterization of a tobamovirus from flowering dogwood in Tennessee.
Plant Dis.
73:174-176.
|
| 36.
|
Smith, P. R.,
R. N. Campbell, and P. R. Fry.
1969.
Root discharge and soil survival of viruses.
Phytopathology
59:1678-1687.
|
| 37.
|
Sobsey, M. D., and P. A. Shields.
1987.
Survival and transport of viruses in soils: model studies, p. 155-177. In
V. C. Rao, and J. L. Melnick (ed.), Human viruses in sediments, sludges and soils.
CRC Press, Inc., Boca Raton, Fla.
|
| 38.
|
Vaughn, J. M., and E. F. Landry.
1983.
Viruses in soils and groundwaters, p. 163-210. In
G. Berg (ed.), Viral pollution of the environment.
CRC Press, Inc., Boca Raton, Fla.
|
| 39.
|
Wait, D. A., and M. D. Sobsey.
1983.
Method for recovery of enteric viruses from estuarine sediments with chaotrophic agents.
Appl. Environ. Microbiol.
46:379-385[Abstract/Free Full Text].
|
Appl Environ Microbiol, April 1998, p. 1430-1435, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
