AN ABSTRACT OF THE THESIS OF
Jonathan L. Whitworth for the degree of Doctor of Philosophy in Crop
and Soil Science presented on April 26, 1993.
Title: Monitoring Potato Leafroll Virus Movement in Differentially Aged
Potato (Solanum tuberosum L.) Plants with an Immunosorbent Direct
Tissue Blotting Assay.
Abstract approved:
Redacted for Privacy
(Alvin R. Mosley)
(Gary L i eed)
Potato leafroll virus (PLRV) causes yield and quality losses in
potato.
PLRV is identified by plant symptoms and serological tests
such as an enzyme-linked immunosorbent assay (ELISA).
A similar
serological test, direct tissue blotting assay (DTBA), was used to
detect and monitor PLRV movement in field-inoculated Russet Burbank
plants and plant tissues from Russet Burbank and Russet Norkotah seed
tubers submitted by growers for winter certification tests.
DTBA was as accurate as ELISA and easier to use for detecting
tuber-perpetuated PLRV in stems and petioles of plants grown from
grower-submitted seed tubers.
ELISA detected twice as many PLRV
positives as DTBA in leaflet tests.
DTBA detected PLRV in tuber tissue
but results matched ELISA in only 74% or less of the samples.
Results
of DTBA tuber tests were sometimes difficult to interpret while stem
and petiole results were distinct and unambiguous.
As inoculations were delayed later in the season and as plants
matured, PLRV infection levels decreased sharply, most often within a
two week period in early July.
In same-age plants inoculated 43 days
after planting but 18 days apart, early inoculation produced higher
PLRV levels.
Conversely, when same-age plants were inoculated 62 days
after planting but 19 days apart, late inoculation produced higher PLRV
levels.
This discrepancy is not fully understood, but larger tuber
size at the later inoculation probably produced a stronger sink for
source-to-sink translocation of nutrients and phloem-limited viruses.
Results of DTBA winter grow-out tests of summer-infected tubers
approximated those of ELISA and visual inspections.
Indirect DTBA
testing of tubers utilizing stem and petiole tissues from winter growout plants detected more PLRV than directly testing tuber tissue 21
days post inoculation in summer.
DTBA detected current season
(primary) PLRV less reliably than secondary (tuber-borne) PLRV, similar
to reported ELISA results.
PLRV infection increased tuber numbers but decreased size.
Size
reduction was most evident in plants infected early in the season.
Average tuber size in healthy plots was always larger than the average
tuber size in infected plots.
Within an infected plant, small tubers
tended to be infected less often than large tubers.
Monitoring Potato Leafroll Virus Movement in Differentially Aged Potato
(Solarium tuberosum L.) Plants with an Immunosorbent Direct Tissue
Blotting Assay.
by
Jonathan L. Whitworth
A THESIS
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Doctor of Philosophy
Completed April 26, 1993
Commencement June 1993
APPROVED:
Redacted for Privacy
A vin R. Mosl
Gary L. R
Professors of Crop and So 1 Science in charge
major
Redacted for Privacy
Head of Department of Crop and Soil Science
Redacted for Privacy
Dean of Graduate
Date thesis is presented
Typed by:
April 26, 1993
Jonathan L. Whitworth
Acknowledgements
This research and my graduate studies would not have been
possible without the help and guidance of Dr. Alvin Mosley and Dr. Gary
Reed.
Their assistance in funding and formulating ideas helped
tremendously.
Dr. Mosley always encouraged me in my studies and
research.
I appreciate the help and advice received from Dr. Harold Toba
and Mr. Lee Fox.
Their generous efforts and knowledge of aphid biology
made it possible for me to do this work.
They supplied aphids for the
field inoculations and were always available to answer questions.
I am indebted to Andrew Jensen, Ph.D. candidate in entomology,
for counting and identifying aphids. collected from yellow pan traps in
the plots.
Before I finish this, I need to acknowledge my wife's gargantuan
effort towards this Ph.D.
Holly provided a sane refuge from sometimes
hectic days and was a good listener and counselor.
Her efforts to
manage home and family may not show up in this printed work, but they
contributed greatly to the quality of it.
Table of Contents
CHAPTER 1
Literature Review
1
Problems Caused by Potato Leafroll Virus
Development of Mature Plant Resistance and Variability in Symptom
Expression
Potato Leafroll Virus Transmission and Translocation
Methods for Detecting Potato Leafroll Virus
Summary of Literature
Literature Cited
2
2
5
6
12
14
CHAPTER 2
Detection of Potato Leafroll Virus by Visual Inspection, Direct Tissue
Blotting and ELISA Techniques.
17
Abstract
Introduction
Methods and Materials
Plant tissues
DTBA methods
ELISA methods
Results
Discussion
17
18
19
19
19
20
21
21
Contributions of Authors
Literature Cited
28
29
CHAPTER 3
Translocation and Detection of Potato Leafroll Virus in Field-Grown
Russet Burbank Plants.
30
Abstract
30
Introduction
32
Methods and Materials
34
Field plot layout and seed preparation
34
Crop production practices
34
Inoculation of field-grown plants with viruliferous aphids
35
Sampling of plants
37
Grow-out tests for PLRV confirmation
37
Inoculation and detection of PLRV in tissue culture-derived
plantlets
38
Statistical analyses
39
Results
40
Plant age and inoculation date effects
40
Reliability of DTBA for detecting PLRV translocation in
field-grown plants
40
Reliability of DTBA for detecting PLRV in tissue culture derived
plantlets
41
PLRV effects on yield
42
Discussion
43
Summary and Conclusions
47
Literature Cited
62
Table of Contents, continued
General Conclusions
Bibliography
63
65
List of Figures
Direct tissue blotting analysis of leaflets (A) healthy
Figure 2.1.
24
leaflet; (B) PLRV-infected leaflet. Magnification 20X.
.
.
.
Stem blots from direct tissue blotting analysis (A)
Figure 2.2.
healthy stem; (B) PLRV-infected stem. Petiole blots are similar.
25
Magnification 20X.
Tuber blots from direct tissue blotting analysis (A)
Figure 2.3.
healthy eye; (B) PLRV-infected sprout on a seed piece. Each blot
is a core of an eye cut and blotted longitudinally.
26
Magnification 20X.
Whole tuber blot from direct tissue blotting
Figure 2.4.
analysis. PLRV-infected daughter tuber (stem end is
uppermost). PLRV antigens are more concentrated at the
stem end in this newly formed daughter tuber.
Magnification 20X.
27
Percentage of potato leafroll virus positive plants
Figure 3.1.
Summer
inoculated at 33 and 48 days after planting (DAP).
test results for stems and tubers; winter test results for
stems of plants grown from summer-tested tubers.
58
Percentage potato leafroll virus infection of plants
Figure 3.2.
inoculated at similar days after planting (DAP) (similar
Summer
growth stages), but on different calendar dates.
test results based on stems and tubers; winter test results
based on stems of plants grown-out from summer-tested
tubers.
59
Natural aphid populations in Russet Burbank plots at
Figure 3.3.
Aphids were collected weekly from
Corvallis, Oregon.
water-filled yellow pan traps located in each corner of the
field.
60
Processed nitrocellulose membranes showing typical
Figure 3.4.
PLRV reactions from inoculation on June 26, 1991 (33 days
Stem and tuber blots from (A) plant
after planting).
samples 5 days post-inoculation; (B) plant samples 17 days
Controls at the bottom of each membrane
post-inoculation.
are 'A', 'B' PLRV-infected; 'C', 'D' healthy.
61
List of Tables
Table 2.1.
Positive potato leafroll virus (PLRV) tests from
visual inspection, direct-tissue blotting and enzyme-linked
immunosorbent assays
23
Table 3.1. Average number of days post-inoculation required for
detectable potato leafroll virus (PLRV) infections in stems
and tubers
49
Table 3.2.
Percentage of Russet Burbank plants and tubers
infected with potato leafroll virus as determined by the
Direct Tissue Blotting Assay.
50
Table 3.3. Number of potato leafroll virus (PLRV) infected
Russet Burbank tubers as determined by Direct Tissue
Blotting Assay (DTBA) and Enzyme-Linked Immunosorbent Assay
(ELISA).
52
Table 3.4.
Potato leafroll virus (PLRV) infection percentages in
stems and nodal cutting& of Russet Burbank plants
inoculated by viruliferous Myzus persicae.
54
Table 3.5.
Yield components from fall-harvested plants with
different levels of potato leafroll virus (PLRV) and from
different planting dates
55
Table 3.6. Correlation coefficients (r2) for assessing
relationships between percentages of plants with potato
leafroll virus (PLRV) at harvest and yield components.
55
.
Table 3.7.
Average tuber size (g) for infected and healthy
tubers from Russet Burbank plants testing positive for
potato leafroll virus (PLRV) using the direct tissue
blotting assay
56
Table 3.8.
Russet Burbank plant growth parameters at potato
leafroll virus inoculation by viruliferous aphids.
57
Monitoring Potato Leafroll Virus Movement in Differentially Aged Potato
(Solanum tuberosum L.) Plants with an Immunosorbent Direct Tissue
Blotting Assay.
CHAPTER 1
Literature Review
Potato leafroll virus (PLRV) significantly increases the cost of
potato production.
PLRV causes depressed yields and poor to
unacceptable tuber quality in some varieties.
important component in PLRV control.
Seed certification is an
Seed stocks must be kept healthy
by roguing infected plants and the use of insecticides to control aphid
vectors.
Seed tubers infected with potato leafroll virus produce
chlorotic, stunted plants that inefficiently translocate photosynthate
to the tubers.
The following season chronically infected plants from
infected tubers serve as primary PLRV inoculum sources for current
season spread of PLRV by aphid vectors.
Success in aphid control
varies among years and requires careful attention each year.
PLRV is a luteovirus which consists of isometric particles
approximately 24 nm in diameter.
Viruses classified as luteoviruses
are icosahedral particles which cause yellowing in plants and are
generally confined to the phloem of the host plant (Francki, 1985).
PLRV is transmitted to plants in a persistent manner by aphid species.
A persistent virus is carried within an aphid's circulatory system and,
once infected, the aphid carries and can transmit the virus for life.
Myzus persicae (Sulzer), the green peach aphid, appears to be the most
efficient and important vector of PLRV although the potato aphid,
Macrosiphum euphorbiae (Thomas), is also reported to transmit PLRV
(Harrison, 1984; MacKinnon, 1969).
Current season or "primary" symptoms in newly infected plants
appear first as pale upper leaves with leaflets rolled and pinched at
the base.
Young leaves in some varieties become pink to reddish along
the base of the leaf margins.
Secondary symptoms, or "chronic"
symptoms occur only in plants grown from infected tubers.
Chronically
infected plants are stunted with rolled lower leaves which feel stiff
and papery (Peters and Jones, 1981).
Problems Caused by Potato Leafroll Virus
Potato leafroll virus causes two major production problems; loss
of yield and poor tuber quality.
Harper et a/. (1975) showed that
yield losses to PLRV in Russet Burbank can range from 65 to 92 percent,
depending on severity of symptoms.
Tuber quality is reduced by the
development of net necrosis in which PLRV-infected phloem cells become
plugged with callose and die (Golinowski and Garbaczeqska, 1984).
These necrosed areas show up as a dark netted pattern in fried chips or
french fries.
A high incidence of net necrosis is sufficient cause for
rejection of potato lots.
Factors other than PLRV such as planting date, nutrition,
variety, and low-temperature storage have been shown to influence the
development of tuber net necrosis (Manzer et a/., 1982).
In one study,
every tuber clearly showing net necrosis contained PLRV, but only 87%
of the tubers with questionable net necrosis symptoms were infected.
The ratio of net necrosis to PLRV varies from 1:1 to 9:1 in seed lots
of cv. Green Mountain (Folsom and Rich, 1940).
Development of Mature Plant Resistance and Variability in Symptom
Expression
To maintain healthy seed, some seed growing areas require that
vines be killed or potatoes harvested when aphids are first detected
each season (Braber et al., 1982).
A good understanding of the time
interval between aphid feeding and viral infection of tubers is
potentially useful information to seed growers and regulatory agencies.
Research shows that mature plant tissues, as opposed to young plants,
slow viral movement from the point of infection to the tubers.
In
young plants, viruses can move into the tubers within five to ten days
3
of infection (Knutson and Bishop, 1964).
Conversely, as the plants age
the virus moves slower and seldom reaches the tubers in old plants.
The hypothesis has been advanced that as a plant matures the proteinsynthesizing capacity declines as the concentration of necessary cell
components (ribosomes and RNA) decreases.
Since viruses replicate
within infected cells, their synthesis is affected by the physiology of
that cell.
Leaves of older plants may not sufficiently multiply
viruses to permit effective translocation to the tubers (Venekamp and
Beemster, 1980).
Researchers have long sought biochemical indicators which may be
used to predict the onset of "mature plant resistance".
Mature plant
resistance appears to be a well established phenomenon for PVX and PVY.
Mature plant resistance is characterized by some degree of resistance
to viral infection and movement in old plants compared to low
resistance to viral infection and translocation in young plants.
Cellular ribosome and RNA content have been shown to drop dramatically
at about the same time resistance to infection by potato virus X (PVX)
or Y (PVY) increases (Venekamp and Beemster,
1980).
1980; Venekamp et al.,
Braber et a/. (1982) measured peroxidase activity and four
different leaf components only to conclude that mature plant resistance
to PVY" did exist but did not correlate with any of Venekamp and
Beemster's suggested controlling factors.
Venekamp and Beemster also
reported that their data did not exclude the possibility that ribosome
and RNA levels drop long before resistance reaches a level of practical
importance and that a causal relationship between cellular components
and mature plant resistance may therefore be remote.
Typical reported dates for the onset of mature plant resistance
to PVY (strain PVY") range from mid-July to August 1 (Braber et a/.,
1982) in the Netherlands, while almost complete resistant was evident
by the beginning of August in Southern Sweden (strain PVY°) (Sigvald,
1985).
Development of mature plant resistance in these two studies was
4
verified by a sudden drop in the percentages of tubers infected and
occurred within a two-week period (Braber et a/., 1982).
Mature plant resistance to PLRV infection has been mentioned
(Beemster, 1972), but few verifying studies have been done.
Storch and
Manzer (1985) inoculated plants late in the season and determined that
plant age had no effect on infection levels observed.
They also found
that the earliest PLRV infection of tubers occurred within three days
of plant infection.
Plants were inoculated at three day intervals when
they were at least 85 days old (from planting).
The previously
described studies with PVX and PVY used plants inoculated at 30 to 77
days from planting.
It is possible that Storch and Manzer's inoculated
plants were too similar in age for age effects to be detected.
PVX and
PVY may also move at different rates than PLRV.
Not only are older plants less likely to become infected with
PLRV late in the season, but when infection does occur, the progeny
tubers produce plants with very mild symptoms.
PLRV symptoms in cv.
Pentland Crown were easily identified in progeny of plants inoculated
in June or July but not in progeny of plants inoculated in August
(Woodford and Barker, 1986).
Testing of progeny plants with enzyme-
linked immunosorbent assays (ELISA) showed that even though late
inoculations produced progeny with very mild PLRV symptoms, they
contained as much virus as progeny plants with severe symptoms from
earlier infections.
In PLRV-ELISA tests, cv. Maris Piper had
absorbance (A400 values of 1.1-1.25 and cv Pentland Crown had values of
0.45-0.51 while healthy plant A46 values were ca. 0.12.
were obtained from all classes of symptom severity.
These values
The A405 values are
proportional to virus concentration (Clark and Adams, 1977).
Knutson
and Bishop (1964) found that progeny from cv. Russet Burbank plants
inoculated with PLRV early in the season expressed leafroll symptoms
earlier than progeny from plants inoculated later in the season.
Among
plants in this study that did become infected following late season
5
inoculations, the percentage of tubers infected per plant was about
half of that in early season inoculations.
Potato Leafroll Virus Transmission and Translocation
An understanding of factors influencing the movement of PLRV into
and within a crop and the time required for the virus to move into the
tubers is essential for developing virus control programs.
In a
disease-free crop, aphid vectors provide the primary source of
inoculum.
PLRV-infected seed in a planting can also serve as a primary
source of inoculum which is then spread to adjacent plants and fields
by alate (winged) aphids (Cadman and Chambers, 1960; Rogerson and
White, 1969; Wright et al., 1970).
Since aphids are the primary vector for PLRV, it is important to
know when high populations match susceptible stages of plant growth.
Seed growing areas usually do not develop high aphid populations until
later in the growing season since cold temperatures usually reduce
overwintering.
Although older potato plants develop a degree of mature
plant resistance, they can still become infected with viruses late in
the season (Flanders et al, 1990; Sigvald, 1985; Storch and Manzer,
1985).
Early season aphid control seems less urgent to growers in
areas with low early season aphid populations compared to areas with
heavy early season aphid pressures.
Regardless, attention to aphid
populations and their control should be given more emphasis when plants
are young and more susceptible to PLRV infection.
The time lapse between foliar inoculation with PLRV and tuber
infection has been reported to vary from eight days in plants of an
unknown age (Bradley and Ganong, 1953) to three days in plants
inoculated from mid-Aug. to mid-Sept (Storch and Manzer, 1985).
Plant
age (from planting to inoculation), time from inoculation to harvest,
planting date, temperature, and date of inoculation all affect the rate
of leafroll virus movement to the tubers (Cadman and Chambers, 1960;
6
Knutson and Bishop, 1964; Storch and Manzer, 1985).
Storch and
Manzer's study was performed in Maine and used multiple regression
analysis to measure the effect of all of the above factors on the
translocation of PLRV to tubers of Russet Burbank.
Time from
inoculation to harvest and temperature accounted for 43% of the PLRV
observed in Russet Burbank.
Storch and Manzer found no real effect of
plant age on the incidence of leafroll-infected tubers (r2 = 0.003).
However, they inoculated late in the growing season so differences in
plant age were small.
In the Knutson and Bishop (1964) Idaho study,
inoculations during late June and the first two weeks of July produced
a high number of plants with primary symptoms while later inoculations
showed few if any visible symptoms.
Similar results were reported by
Cadman and Chambers (1960) in Scotland.
Results showing higher
infection rates from inoculations in late June to early July and low
rates from late season inoculations corroborate studies discussed
previously concerning mature plant resistance.
Methods for Detecting Potato Leafroll Virus
Studies described above are based on visual plant symptoms and
are sometimes confirmed with grow-out tests of progeny tubers.
In
1977, R. Casper showed that an enzyme-linked immunosorbent assay
(ELISA) could detect PLRV in potato foliage and tubers.
This finding
was an important breakthrough because, up to that point, serological
detection of PLRV was performed with either the Ouchterlony gel-
diffusion test with PLRV-specific antiserum or the Ingel-Lange test to
detect PLRV associated callose in tubers (van Slogteren, 1972; de Bokx,
1972).
Neither of these tests were amenable to rapid screening of
large numbers of samples for certification purposes.
ELISA had
previously been used with a high success rate to detect small amounts
of other viruses in other host plants (Clark, et al., 1977; Clark and
Adams, 1977).
Since the 1977 Casper study ELISA has become a standard
7
test for detecting potato viruses in many programs.
ELISA is also useful for research purposes because
results not
only indicate viral presence, but also the relative
concentration
(Clark and Adams, 1977).
In the ELISA technique, viral antigens attach
to virus-specific immunoglobulins which are coated on
a solid-phase
support system (usually a multi-well polystyrene plate).
Bound
antigens are detected with virus-specific antibodies
conjugated to
alkaline phosphatase. This enzyme is reacted
with p-nitrophenyl
phosphate in a substrate buffer to produce a yellow color.
Light
absorbance values are determined with a photometer and the
value is a
function of, and proportional to, the virus concentration.
ELISA is also used to test PLRV concentration in tubers,
(Casper,
1977; Clarke, et al., 1980; Gugerli, 1980; Hill and
Jackson, 1984;
Tamada and Harrison, 1980b) stems, roots, and leaves
(Casper, 1977).
In most of the tuber studies, the concentration of PLRV
is higher in
the stolon end than the bud end at harvest and after
natural or
artificial dormancy break (Clarke et a/., 1980;
Gugerli, 1980; Tamada
and Harrison, 1980b). The difference in PLRV
concentration between the
stolon and bud ends of tubers is less pronounced after
artificial
dormancy break and a warming period of approximately
two weeks
(Gugerli, 1980).
All three of these papers verify the utility of
ELISA
for detecting PLRV in tubers. Hill and Jackson (1984)
used Gugerli's
extraction method and found that ELISA bud end tests
resulted in a
serious underestimation of the virus when compared to ELISA tests using
foliage from grow-outs of the same tubers.
However, Hill and Jackson
did not compare samples from stolon end tuber tissue.
One study with
contrasting results was done by Weidemann (1988) that
showed higher
concentrations of PLRV present in the bud end. A study of several
tissues by Flanders et a/. (1990) produced the most
accurate ELISA
results with foliage from tuber grow-outs; PLRV
detected by this method
would be tuber-borne (secondary infection) indicating
that the parent
8
plant was infected.
When tubers were harvested 21 to 28 days after
the
plants had been artificially inoculated with PLRV,
testing the foliage
from a primary-infected tuber failed to detect
virus in seven percent
of the infections. Testing similar plants
harvested 7 to 14 days after
inoculation missed 67% of the infections.
Most certification agencies test seed lots by planting
tuber
samples in a greenhouse or in fields in southern
states during the
winter and visually scoring the plants for
virus symptoms. Even though
the concentration of PLRV is greater in the
stolon end than in the bud
end of a tuber, plants from bud end eye pieces
produce the most
accurate results when compared to mid-tuber
or stolon-end eyes (Hoyman,
1962).
Higher metabolic activity in the bud end during
sprouting would
result in increased virus replication making
the bud end the best
tissue to sample for grow-outs and subsequent
virus detection.
Tubers
exhibit apical dominance and sprouts develop
first at the bud end.
When questionable plant symptoms are encountered,
ELISA can be used to
confirm visual results. ELISA is also widely used
to test large sample
numbers for latent viruses such as potato virus X
(Wright et al. 1976),
which are not easily detected by visual observation.
The use of polystyrene solid-phase plates for
antigen binding in
ELISA is common.
Efforts to increase the sensitivity of ELISA
have
utilized other solid support systems.
Attempts have also been made to
simplify existing ELISA techniques to reduce time
requirements so that
more samples can be processed.
Nitrocellulose membranes (NCM) have a high affinity
for proteins
and were first used to detect potato viruses S,
X, and Y by Banttari
and Goodwin (1985).
The term "dot-ELISA" refers to the use of
NCM
instead of polystyrene plates for attachment of
viral antigens.
PLRV
is phloem-limited and its low concentrations
in sap extracts can make
it more difficult to detect in assays than
PVX or PVY which tend to
infect every cell. Smith and Banttari (1987)
found dot-ELISA to be
9
twice as sensitive as polystyrene based ELISA
(multi-well plates) for
detecting PLRV in young potato leaves.
NCM could be pre-coated with
trapping antibodies in order to reduce the time required
to run the
actual assay. Samples on NCM were stained
with fast red TR salts and
produced positive red color reactions (denoting the
presence of viral
antigens); when dried, samples could be stored
as permanent records for
as long as six months. Results were quantified
by measuring the color
reaction with a reflectance densitometer.
In another study using this
technique, tuber and leaf extracts were used in
a dot-ELISA and the
results were equal to ELISA (technique of Clark and
Adams, 1977), but
results were based on a visual assessment which is
more subjective.
The author also applied plant sap from potato
virus S (PVS)-infected
stems directly to the NCM by squeezing them with
forceps and then
pressing the tissue onto the membrane (Weidemann,
1988). The concept
of directly applying plant sap from the cut
surface of intact tissue
was later developed into a very rapid, simple, and
sensitive virus test
by Lin et al. (1990). Other techniques using
plain notebook paper
(Heide and Lange, 1988) or filter paper (Haber and
Knapen, 1989) were
not sensitive enough for luteoviruses or less sensitive
than ELISA.
Another convenience of dot-ELISA on NCM is that the
unprocessed samplespotted NCM can still produce reliable results after 20
weeks of
storage or after being sent through the mail (Lizarraga
and Northcote,
1989).
The dot-ELISA technique includes a pre-treatment
of the NCM with
trapping antibodies and the use of plant sap extracts.
In a more
simple test developed by Lin et al. (1990), intact plant
tissue is cut
and directly blotted onto untreated NCM.
Reagents and reagent
concentrations similar to those for ELISA are used.
Positive results
are indicated by the formation of a purple color reaction
from an
10
NBT/BCIP1 substrate which can be detected
with the naked eye although a
dissecting scope at 20X is helpful. This test is potentially
more
sensitive than other tests which require
a dilution of the sap extract
with buffer before use.
In a polystyrene-based
plate system, 200 it of
sap extract is usually applied.
The limiting factor for NCM is the
amount of antigens present at the cut surface.
Hsu and Lawson (1991)
showed that dot-ELISA (on NCM) was eight times
more sensitive than
polystyrene-based ELISA. Tests with viruses in
ELISA plates indicate
that only about 25 to 37% of antigens applied
to each well were
actually trapped by the coating antibodies.
NCM's are estimated to
have a protein binding capacity of 80 pl/cm2 which
permits the
absorption of all viral antigens in a sample
solution (Gershoni and
Palade, 1982). Precise comparisons of
sensitivity between NCM-based
and other systems is difficult because different
plant tissue
preparations are used.
Since intact tissue is used with the direct
tissue blotting assay (DTBA), there is no
way to do dilution endpoint
studies to test sensitivity.
Bravo-Almonacid, et al. (1992) used DTBA
to detect PVX and PVY in potato. They estimated the
sensitivity of
DTBA to be about 0.25 ng virions per 10 mg tissue,
while their current
double antibody sandwich ELISA was sensitive to
approximately 5 ng.
To
obtain these figures, they made the signal
at the limit of detection
from a virus standard (with a known amount of virions)
equal to the
limit of detection for infected tissue.
They stated that strict
quantitative comparisons at the level of tissue
samples were difficult
to establish.
The direct tissue blotting on NCM assay method
was shown to be
effective on a number of viruses, including potyvirus
and potexvirus
groups (Lin et al., 1990).
Its use on the luteovirus barley yellow
dwarf was successful in detecting the pathogen, but
results were
'
nitro-blue
phosphate tetrazolium/5-bromo-4-chloro-3-indolyl-
11
inconsistent.
Barley yellow dwarf antigens were not detected
in every
cross section blot from infected tissue, but all blots
from healthy
leaves were negative. The authors used leaf
tissue for the tests and
speculated that inconsistent results were probably
caused by the uneven
distribution of antigen in the phloem of infected
leaves.
The tissue
blots showed that barley yellow dwarf virus
was confined to the phloem.
A comparison of DTBA with ELISA for detection
of potato viruses showed
about equal precision for potato viruses X
and Y, while S and A were
detected more often by DTBA, and PLRV was
detected more often by ELISA
(Samson et a/., 1993). DTBA has the advantage of
fewer steps than dotELISA or polystyrene based ELISA, and is simpler
because no prepreparation of NCM or plant tissue is required.
Viral antigens are
also localized in plant tissue making DTBA
a valuable study method for
translocation of viruses (Lin et a/., 1990).
The disadvantage of DTBA
is that virus concentrations
cannot presently be quantified.
The DTBA is extremely suitable for seed
certification, foundation
seed, and variety development programs where
a large number of samples
are screened for the presence of viruses.
It permits the use of plant
tissues directly in the field or in places other
than well equipped
laboratories. Tissue blotted NCM
can be sent through the mail for
processing by another lab or can be processed with
a minimal number of
reagents in a basic laboratory.
Materials required include a
dissecting microscope (as an aid in interpretation),
a supply of razor
blades, and a refrigerator for storing reagents.
12
Summary of Literature
Potato leafroll virus reduces yield and quality in susceptible
cultivars.
PLRV is vectored by aphids (most efficiently by Myzus
persicae).
The use of healthy seed stock and thorough control of
aphids provide the best means of commercial control.
The production of
healthy seed is the responsibility of seed growers and certification
agencies.
Certification personnel inspect fields at least twice during
the growing season and conduct winter grow-out tests to detect latent
viruses (PVX) and other viruses introduced late in the season.
Late
season PLRV infections usually do not show any visual plant symptoms,
but some tubers in a hill may still become infected.
Potato plants develop a degree of "mature plant resistance" to
virus infection late in the season.
Late season inoculation of plants
(mid-July into August) in the United States results in a greatly
reduced infection percentage (Knutson and Bishop, 1964), although the
virus may move more rapidly to the tubers if infection does occur
(Storch and Manzer, 1985).
Detection of current season leafroll (primary infection) is
unreliable with enzyme-linked immunosorbent assay (ELISA) techniques
because viral antigens, especially PLRV antigens, may be unevenly
distributed in the plant.
Tuber borne viruses (secondary infection)
produce plants that are uniformly infected and produce the most
reliable ELISA results (Flanders et al., 1990).
ELISA techniques are
easily mechanized for processing large sample numbers, but much time is
required for sample preparation (plant sap extraction and dilution, and
loading samples into multi-well assay plates).
For simple detection of
the presence of a virus, the direct tissue blotting assay (DTBA)
offers a distinct advantage.
Freshly cut tissues are blotted directly
onto untreated nitrocellulose membranes without any special tissue
preparation.
This method is simpler, faster, and as sensitive, or more
sensitive, than ELISA (Bravo-Almonacid et al, 1992; Hsu and Lawson,
13
1991).
DTBA offers certification personnel and growers a simple,
valuable tool for use in screening seed stocks for viruses.
DTBA is as
precise as ELISA for detecting potato viruses X, Y (Samson
et al.,
1993), and leafroll (Whitworth et al., 1993).
A better understanding of the time required for PLRV to move into
the tubers of different-aged plants and the percentage of
those tubers
which become infected can be gained by the use of DTBA.
This
understanding would help in determining when to spray plants to control
aphids and when plants are most likely to be susceptible
to infection.
14
Literature cited
Banttari, E.E. and P.H. Goodwin. 1985.
Detection of potato viruses S,
X, and Y by enzyme-linked immunosorbent assay on nitrocellulose
membranes (dot-ELISA). Plant Disease 69:202-205.
Beemster, A.B.R.
1972.
Virus translocation in potato plants and
mature plant resistance.
In Viruses of potatoes and seed-potato
production. (ed.) deBokx J.A.
Cen. Agric. Pub. Wageningen.
Braber, J.M., C.B. Bus, and A. Schepers.
1982.
Changes in leaf
components and peroxidase activity of potato plants (cv. Bintje)
in relation to mature-plant resistance to PVYN. Potato Research
25:141-153.
Bradley, R.H.E. and R.Y. Ganong.
1953.
Note on the time potato leaf
roll virus takes to pass from aphids on the leaves into the
developing tubers.
Can. J. Bot. 31:143-4.
Bravo-Almonacid F., L. Haim, and A. Mentaberry.
1992.
Rapid
immunological detection of potato viruses in plant tissue
squashes.
Plant Disease 76:574-578.
Cadman, C.H. and J. Chambers. 1960.
Factors affecting the spread of
aphid-borne viruses in potato in Eastern Scotland. Ann. Appl.
Biol. 48:729-738.
Clark, M.F. and A.N. Adams.
1977.
Characteristics of the microplate
method of enzyme-linked immunosorbent assay for the detection of
plant viruses. J. Gen. Virol. 34:475-483.
Clark, M.F., A.N. Adams, J.M. Thresh, and R. Casper, 1977. The
detection of plum pox and other viruses in woody plants by
enzyme-linked immunosorbent assay (ELISA).
Acta Hortic. 67:5157.
Casper, R.
1977.
Detection of potato leafroll virus in potato and
Physalis floridana by Enzyme-linked immunosorbent assay (ELISA).
Phytopath. 90:364-8.
de Bokx, J.A.
1972.
Histology, cytological and biochemical methods.
p. 111-114. In J.A. de Bokx (ed.)
Viruses of potatoes and seedpotato production. Center for Agricultural Publishing and
Documentation, Wageningen, Netherlands.
Flanders, K.L., D.W. Ragsdale, and E.B. Radcliffe.
1990.
Use of
enzyme-linked immunosorbent assay to detect potato leafroll virus
in field grown potato, cv. Russet Burbank.
Amer. Potato J.
67:589-602.
Folsom, D. and A.E. Rich.
1940.
Potato tuber net-necrosis and stemend browning studies in Maine. Phytopath. 30:313-322.
Francki, R.I.B.
Florida.
1985.
Atlas of plant viruses.
CRC Press, Coca Raton,
Gershoni, J.M. and G.E. Palade.
1982.
Electrophoretic transfer of
proteins from sodium dodecyl sulphate polyacrylamide gels to a
positively charged membrane filter.
Anal. Biochem. 124:396-405.
15
Golinowski W. and G. Garbaczewska. 1984. Process of necrosis
formation in potato plants infected with potato leaf roll virus.
Zeszyty Problemowe Postepow Nauk Rolniczych 298:45-53.
Gugerli, P. 1980.
Potato leafroll virus in vascular region of potato
tubers examined by ELISA. Potato Res. 23:137-41.
Haber, S. and H. Knapen.
1989.
Filter paper sero-assay (FiPSA):
A
rapid, sensitive technique for sero-diagnosis of plant viruses.
Can. J. Plant Path. 11:109-113.
Harper, F.R., G.A. Nelson, and U.J. Pittman. 1975.
Relationship
between leafroll symptoms and yield in Netted Gem potato.
Phytopath. 65:1242-4.
Harrison, B.D. 1984. Potato leafroll virus.
Plant Viruses. No. 291. 6 pgs.
CMI/AAB Description of
Heide, M. and L. Lange.
1988.
Detection of potato leafroll virus and
potato viruses M, S, X, and Y by dot immunobinding on plain
paper.
Potato Res. 31:367-73.
Hill, S.A. and E.A. Jackson.
1984.
An investigation of the
reliability of ELISA as a practical test for detecting PLRV and
PVY in tubers.
Plant Path. 33:21-26.
Hoyman, W.G. 1962.
Importance of tuber eye position when indexing for
the leafroll virus. Amer. Potato J. 39:439-43.
Hsu, H.T. and R.H. Lawson.
1991. Direct tissue blotting for detection
of tomato spotted wilt virus in impatiens. Plant Disease 75:29295.
Knutson, K.W. and G.W. Bishop.
1964.
Potato leafroll virus - Effect
of date of inoculation on percent infection and symptom
expression. Amer. Potato J. 41:227-238.
Lin, M.S., Y.H. Hsu, and H.T. Hsu.
1990.
Immunological detection of
plant viruses and a mycoplasma-like organism by direct tissue
blotting on nitrocellulose membranes. Phytopath. 80:824-28.
Lizarraga, C. and E.N. Fernandez-Northcote. 1989. Detection of potato
viruses X and Y in sap extracts by a ;modified indirect enzymelinked immunosorbent assay on nitrocellulose membranes (NCMELISA).
Plant Dis. 73:11-14.
MacKinnon, J.P.
1969.
Transmission of leaf roll virus from potato to
potato by a green clone of Macrosiphum euphorbiae. Amer. Potato
J. 46:23-24.
Manzer, F.E., D.C. Merriam, R.H. Storch, and GW Simpson.
1982.
Effect
of time of inoculation with potato leafroll virus on development
of net necrosis and stem-end browning in potato tubers. Amer.
Potato J. 59:337-49.
Peters, D. and R.A.C. Jones.
1981.
Potato leafroll virus In W.J.
Hooker (ed.) Compendium of potato diseases. 125 pgs. The
American Phytopathology Society. APS Press.
16
Rogerson, J.P. and J.H. White.
1969. Aphids and the spread of leaf
roll virus in seed potatoes in the North Riding of Yorkshire,
1964-66.
Plant Path.
18:156-63.
Samson, R.G., T.C. Allen, and J.L. Whitworth.
1993.
Evaluation of
direct tissue blotting to detect potato viruses.
Amer. Potato J.
(in press).
Sigvald, R.
1985.
Mature-plant resistance of potato plants against
potato virus Y° (PVY°).
Potato Research 28:1355-143.
Smith, F.D. and E.E. Banttari.
1987.
Dot-ELISA on nitrocellulose
membranes for detection of potato leafroll virus. Plant Dis.
71:795-99.
Storch, R.H. and F.E. Manzer.
1985.
Effect of time and date of
inoculation, plant age, and temperature on translocation of
potato leafroll virus into potato tubers.
Am Potato J 62:137-43.
Tamada, T. and B.D. Harrison.
1980a.
Factors affecting detection of
PLRV in potato foliage by ELISA. Ann. Applied Biol.
95:209-19.
Tamada, T. and B.D. Harrison.
1980b. Application of ELISA to
detection of PLRV in potato tubers. Ann. Applied Biol.
96:6778.
van Slogteren, D.H.M., 1972.
Serology. p.87-101. In J.A. de Bokx
(ed.)
Viruses of potatoes and seed-potato production. Center
for Agricultural Publishing and Documentation, Wageningen,
Netherlands.
Venekamp, J.H., A. Schepers, and C.B. Bus.
1980.
Mature plant
resistance of potato against some virus diseases.
III.
Mature
plant resistance against potato virus YN, indicated by decrease
in ribosome-content in ageing potato plants under field
conditions.
Neth. J. Plant Path. 86:301-309.
Venekamp, J.H. and A.B.R. Beemster. 1980. Mature plant resistance of
potato against some virus diseases.
II.
Mature plant resistance
and the influence of temperature on the ribosome and RNA content
in leaves.
Neth. J. Plant Path. 86:11-16.
Weidemann, H.L.
1988.
Rapid detection of potato viruses by dot-ELISA.
Potato Res. 31:485-92.
Whitworth, J.L., R.B. Samson, T.C. Allen, and A.R. Mosley. 1993.
Detection of potato leafroll virus by visual inspection, direct
tissue blotting and ELISA techniques.
Amer. Potato J. (in
press).
Woodford, J.A.T. and H. Barker.
1986.
Problems of detecting potato
leafroll virus following late infection. Aspects of Applied
Biol.
13:325-31.
Wright, H.S., J.E. Cochran, F.E. Manzer, and J. Munro. 1976. Report
of committee on PVX testing. Amer. Potato J. 53:333-335.
Wright, N.S., H.R. MacCarthy, and A.R. Forbes. 1970.
Epidemiology of
potato leafroll virus in the Fraser River delta of British
Columbia. Amer. Potato J. 47:1-8.
17
CHAPTER 2
Detection of Potato Leafroll Virus by Visual Inspection, Direct Tissue
Blotting and ELISA Techniques.
Abstract
Visibly infected and healthy Russet Burbank and Russet Norkotah
plants grown from tubers submitted by seed growers for winter
greenhouse grow-out trials were tested for potato leafroll virus (PLRV)
by visual examination, enzyme-linked immunosorbent assay (ELISA) and
direct tissue-blotting assay (DTBA).
Each plant was sectioned into
leaflet, petiole, stem, root, seed piece, and tuber portions.
Tissues
were blotted on a nitrocellulose membrane for DTBA and then homogenized
for use in a double-antibody sandwich ELISA system.
Agreement between
the two serological detection methods and with visual readings was high
for petioles and stems, but lower for leaflet, tuber and root tissues.
Comparison of DTBA with ELISA and with visual plant symptoms suggest
that DTBA can be used with the same accuracy as ELISA for detecting
PLRV in stems and petioles.
18
Introduction
Potato leafroll virus (PLRV), a luteovirus, can reduce yields,
Leafroll-infected tubers of Russet
tuber size and internal quality.
Burbank and certain other cultivars are highly susceptible to the
development of net necrosis.
Most of the Pacific Northwest Russet
and
Burbank crop is processed into frozen french fries from storage
high levels of net necrosis make the tubers unacceptable for
processing.
Accurate detection of PLRV is important for producing
healthy seed and controlling net necrosis in commercial crops.
The direct tissue blotting method developed by Lin, et al. (1990)
number of viruses,
uses an indirect immunological procedure to detect a
including an RPV isolate of barley yellow dwarf (BYDV) luteovirus.
Detection of BYDV-RPV antigens in leaf sections with this method was
not consistent in that antigens were not detected in blots from all
cross sections.
Tissue blots from healthy barley leaves did not show
BYDV antigen-specific labels.
Lin, et al. surmised that the
inconsistency of the test in infected tissue was due to the uneven
distribution of BYDV antigens.
Another study found that enzyme-linked
immunosorbent assay (ELISA) detected more PLRV positives in potato leaf
samples than did the direct tissue blotting assay (DTBA) (Samson et
al., 1993).
This study was designed to determine the extent to which results
diagnosis for
of DTBA and ELISA agree with each other and with visual
detecting PLRV in various potato tissues.
19
Methods and Materials
Plant tissues
Plants of Russet Burbank and Russet Norkotah were grown from
excised eyes of tubers submitted for winter certification tests.
Plants with and without visible PLRV symptoms were sectioned into
leaflets, petiole, stem, root, seed piece, and tuber tissues.
Tissues
were blotted on nitrocellulose membrane for DTBA analysis and then
homogenized for use in a double antibody-sandwich ELISA system.
For
each leaf blot, three leaflets were rolled tightly together, cut
transversely with a sterilized razor blade, and pressed onto the
nitrocellulose membrane.
Stems and petioles were blotted singly.
Roots from individual plants were combined,
folded in half, cut
transversely, and blotted. Cores six mm in diameter were removed from
sprouted eyes of seed pieces, sectioned longitudinally and blotted.
Stem-end cores of daughter tubers were sectioned longitudinally and
blotted.
Tubers less than one cm in diameter were divided
longitudinally and blotted directly without coring.
Because petioles
appeared to be especially well suited for DTBA, an additional 241
petiole samples were tested to further compare the reliability of DTBA
and ELISA sensitivity for detecting PLRV in petioles.
DTBA methods
The blotted membranes were processed in
petri dishes using a
direct immunological test modified from Lin et al (1990).
Membranes
were first immersed for 60 min. at room temperature in a blocking
buffer made of non-fat dry milk (1 g/l) in phosphate-buffered saline
with Tween (PBS-Tween) (0.14 M NaC1, 0.01 M Na2HPO4, 1.5 mM KH2PO4, 2.7
mM KC1, 3.1 mM NaN3, and 0.05% Tween-20, adjusted to pH 7.4).
Membranes were then incubated overnight at 4° C in polyclonal antiserum
(1:800 dilution) conjugated to alkaline phosphatase (Boehringer
Mannheim, Indianapolis, IN).
The same batch of antiserum was used to
20
process both DTBA and ELISA tests.
Membranes were then rinsed for 10-
15 min three times in PBS-Tween and incubated for about 40 min, or
until sufficiently developed, in substrate buffer containing 0.1 M
Tris, 0.1 M NaC1, and 5 mM MgC12 at pH 9.5.
Nitroblue tetrazolium (14
mg/40 ml buffer) and 5-bromo-4-chloro-3-indoly1 phosphate
(7 mg/40 ml
buffer) were added to the stock substrate buffer solution just before
use.
Blots from healthy and PLRV-infected control plants were added to
each membrane.
Each membrane was processed in the same petri dish
throughout the procedure, with approximately 10 ml of reagent added and
removed at each step.
DTBA tests were considered positive when
examination of the blots under a 20X dissecting'scope showed purpling
in phloem tissues.
Since DTBA results are subjective, readings were
considered positive only when two researchers independently examining
the same tissue blots agreed.
ELISA methods
Tissue extracts (1-2 gm tissue homogenized in 3 ml buffer) were
Each
placed into microplates coated with a PLRV-specific antiserum.
plate contained four wells with a known PLRV-infected control, two
wells with sample buffer only, six wells with a known healthy control,
and two wells for each unknown sample.
The same antiserum with
conjugate described for DTBA was used.
ELISA procedures followed
methods described by Clark and Adams (1977).
ELISA results were
considered to be positive when absorbance readings with a Bio-Tek
photometer set at 405 nm exceeded both 0.100 OD and a healthy plant
mean absorbance reading by four standard deviations.
DTBA and ELISA results were considered to be in agreement only
when both researcher's DTBA readings and ELISA results matched.
were examined and photographed through a dissecting scope at 20X
magnification.
Blots
21
Results
ELISA detected more PLRV positives in leaflets than DTBA,
especially in upper leaves (Table 2.1).
Interpretation of DTBA leaflet
blots was visually difficult because phloem bundles were unevenly
dispersed (Figure 2.1).
Stems and petioles provided the closest agreement between the two
serological assays and were easiest to interpret for DTBA (Table 2.1).
Roots were difficult to test with DTBA because of limited tissue
volume.
Agreement between DTBA and ELISA tests on tuber tissue was better
for daughter tubers than for seed pieces, but relatively poor for both.
.Interpretation of DTBA tests was less precise for tubers than for stems
and petioles.
tests.
DTBA and ELISA results did not agree for 15 of 41 tuber
Seven of the conflicting tests were DTBA positive and ELISA
negative, one showed the reverse pattern, and interpretation of the
remaining seven produced conflicting DTBA results.
Figure 2.2
illustrates the distribution of PLRV antigens in tuber tissues.
Stem and petiole readings matched results of previous visual
DTBA and ELISA
foliar inspections better than those of other tissues.
stem tests matched visual positives exactly, while DTBA petiole tests
missed one positive detected by both ELISA and visual examination.
Discussion
Concentrated phloem cells in stems and petioles produce DTBA
results equally as reliable as those of ELISA.
Leafroll is a phloem-
limited virus (Harrison, 1984) and infected stem and petiole phloem
cells turn purple during DTBA, making results unambiguous and easy to
interpret.
DTBA petiole analysis is especially useful for non-
destructively sampling large numbers of plants for PLRV.
Agreement between DTBA and ELISA for leaflets was poor. Visual
PLRV-positives for leaflets were more closely matched by ELISA than by
22
DTBA.
The lower number of leaflet positives with DTBA was most likely
due to the absence of concentrated phloem bundles compared to stems and
petioles.
DTBA tissue blots detect virus antigens only in phloem cells
at the surface of the cut; thus, results are inconsistent for leaflets
and other tissues with scattered phloem concentrations.
ELISA tests used homogenized tissue extracts which potentially
represent all phloem cells and any associated antigens, therefore ELISA
is superior to DTBA for testing leaflets.
A previous study showed the
most reliable tissue for detecting PLRV by ELISA to be plant foliage
grown from infected tubers (Flanders et a/., 1990).
DTBA can be used to determine the distribution of PLRV in potato
tissues such as sprouts and new tubers (Figure 2.2).
The fact that
tissue blots detect virus antigens present on a cut surface at any
chosen location makes DTBA useful for monitoring PLRV movement in a
plant or tuber (Figures 2.3 and 2.4).
DTBA cannot quantify virus titre.
However, DTBA can be used by
seed growers and certification agencies to quickly detect and/or
confirm the presence of PLRV using stems or petioles.
DTBA should also
be useful in detecting other potato viruses (Samson et a/., 1993).
23
Table 2.1. Positive potato leaf roll virus (PLRV) tests from visual
inspection, direct-tissue blotting and enzyme-linked immunosorbent
assays.
PLRV-positives by:
% agreement between
No. of
samples
ELISA
DTBA
Visual'
Assays'
Upper
leaves
33
24
12
23
55
67
Lower
leaves
12
8
6
8
58
58
Petioles
33
23
22
23
97
100
Stems
33
23
23
23
97
100
Roots
30
13
7
20
17
49
Seed
pieces
18
8
10
11
61
73
Daughter
tubers
23
17
16
17
74
87
Petioles
2414
24
26
NA5
NA
NA
Tissue
Evaluators'
1 Plants with visible foliar symptoms prior to tissue sampling
2 Percentage = (No. of tests DTBA=ELISA / total tests) X 100; includes
both positive and negative results
3 Percentage = (number of tests for which both evaluators agree / total
number of tests) X 100
4 Samples submitted to T.C. Allen's lab for PLRV testing
5 Data not available
24
Figure 2.1. Direct tissue blotting analysis of leaflets (A) healthy
leaflet; (B) PLRV-infected leaflet. Magnification 20X.
25
Stem blots from direct tissue blotting analysis (A)
Figure 2.2.
healthy stem; (B) PLRV-infected stem. Petiole blots are similar.
Magnification 20X.
26
Tuber blots from direct tissue blotting analysis (A)
Figure 2.3,
Each blot is a
healthy eye; (B) PLRV-infected sprout on a seed piece.
core of an eye cut and blotted longitudinally. Magnification 20X.
27
Whole tuber blot from direct tissue blotting analysis.
Figure 2.4.
PLRV antigens
PLRV-infected daughter tuber (stem end is uppermost).
are more concentrated at the stem end in this newly formed daughter
Magnification 20X.
tuber.
28
Contributions of Authors
The manuscript for this chapter was accepted for publication by
the American Potato Journal Nov. 2, 1992 and is currently in press
(Vol. 70).
Authors and their contributions are:
Jonathan L. Whitworth, Graduate Student;
conducted experiments,
analyzed results, prepared preliminary and final drafts.
Richard G. Samson, Research Assistant;
provided assistance, reagents,
and visual interpretation of direct tissue blotting results.
Thomas C. Allen, Virologist;
provided laboratory space, advice and
encouragement.
Alvin R. Mosley, Agronomist; provided editing skills, advice, funding,
and encouragement.
This work was supported in part by the UDSA/CSRS and the Oregon
Potato Commission and will be published as Oregon State University
Agricultural Experiment Station technical paper no. 9901.
29
Literature cited
Clark, M.F. and A.N. Adams.
1977.
Characteristics of the microplate
method of enzyme-linked immunosorbent assay for the detection of
plant viruses. J Gen Virol 34:475-483.
Flanders, K.L., D.W. Ragsdale, and E.B. Radcliffe.
1990.
Use of
enzyme-linked immunosorbent assay to detect potato leafroll virus
in field grown potato, cv. RusSet Burbank. Amer. Potato J.
67:589-602.
Harrison, B.D.
1984.
Potato leafroll virus.
Plant Viruses. No. 291.
6 pgs.
CMI/AAB Descriptions of
Lin, N.S., Y.H. Hsu, and H.T. Hsu. 1990.
Immunological detection of
plant viruses and a mycoplasma-like organism by direct tissue
blotting on nitrocellulose membranes. Phytopathology 80:824828.
Samson, R.G., T.C. Allen, and J.L. Whitworth. 1993.
Evaluation of
direct tissue blotting to detet potato viruses. Amer. Potato J.
70:257-265.
30
CHAPTER 3
Translocation and Detection of Potato Leafroll Virus in Field-Grown
Russet Burbank Plants.
Abstract
Potato leafroll virus (PLRV) is transmitted by aphids and reduces
potato tuber yield and quality.
PLRV is best controlled by using
healthy seed and insecticides to minimize aphid populations.
The time
required for PLRV to move from foliar infection sites to tubers is
affected by plant age and the seasonal time of inoculation by
viruliferous aphids.
plants is poor.
Visual symptom expression in late season infected
Primary PLRV infections from late season inoculation
are best confirmed by winter grow-out tests, while tuber-borne
(secondary) PLRV can be reliably detected by serological methods such
as the enzyme-linked immunosorbent assay (ELISA).
The direct tissue
blotting assay (DTBA), a similar serological test, is as reliable as
ELISA for detecting secondary PLRV and may be potentially more
sensitive because undiluted plant sap is used in the test.
Russet Burbank seed potatoes were planted at one date in 1991 and
two dates in 1992 to provide plants of different maturities which were
then inoculated with PLRV by introducing viruliferous aphids in late
June, mid-July, and early August.
Aphids were allowed to feed for two
to three days before plots were sprayed with an insecticide.
Plants
and tubers were tested for PLRV with DTBA at periodic intervals after
inoculation.
Tubers were indexed and grown-out in the greenhouse the
following winter to confirm virus presence and results of serological
tests.
Plant age affected plant infection percentages more consistently
than did the actual inoculation date.
As plants aged, fewer became
infected with each succeeding inoculation.
When young same age plants
(-43 days from planting) were inoculated at different dates, the early
inoculation resulted in the highest percentage of infected plants.
31
Conversely, for unknown reasons, when old same age plants (-62 days
from planting) were inoculated at different dates, the late inoculation
resulted in the highest percentage of infected plants.
In all cases,
early inoculation of young plants resulted in the highest infection
percentages.
DTBA is best used to detect secondary PLRV present in the foliage
of plants grown from infected tubers.
DTBA is less reliable for
detecting primary (current season) PLRV.
Visual evaluation, ELISA, and
DTBA were equally effective for detecting secondary PLRV in winter test
plants.
32
Introduction
Symptom expression from current season (primary) PLRV infection
of potato plants is variable and may be absent when infection occurs
late in the season.
Potato plants develop some degree of resistance to
viruses as they mature.
Tuber infection percentages typically decrease
as plant infection is delayed until later in the season and large
tubers tend to be infected more often than small tubers (Knutson and
Bishop, 1964).
The use of larger tubers for grow-out detection of PLRV
would probably improve the accuracy of readings.
An understanding of
relative plant susceptibility to PLRV and current aphid populations
would be useful in controlling the spread of PLRV.
If aphids are
present when plants are young and more susceptible to infection, then
preventive insecticide sprays should be applied more frequently and
with greater diligence compared to later in the season when plant
susceptibility is low.
A better understanding of the time required for
PLRV to move from infection sites to tubers of different aged plants is
needed to optimize vine killing and harvest dates.
A quick and reliable serological test for both current season and
tuber-borne PLRV would be useful for research as well as seed
certification purposes.
The enzyme-linked immunosorbent assay (ELISA)
is less reliable for detecting current season (primary) PLRV than
tuber-borne (secondary) PLRV (Flanders et a/.,
1990).
The recently
developed direct tissue blotting assay (DTBA) is a relatively simple,
but accurate serological test.
It is as reliable as ELISA for
detecting tuber-borne (secondary) PLRV in cvs. Russet Burbank and
Russet Norkotah plants (see Chap. 2).
DTBA utilizes the same enzyme-
linked antibody tag detecting system used in ELISA.
However, intact
tissue is blotted directly onto a nitrocellulose membrane in DTBA,
while tissues are homogenized, diluted, and suspended in a buffer for
ELISA.
DTBA has been reported to be relatively more sensitive than
ELISA for detecting potato viruses X and Y (Bravo-Almonacid et a/.,
33
1992), but direct comparison between the two methods is difficult
because of the different tissue preparations required.
Because
undiluted plant sap is used to apply viruses to nitrocellulose in DTBA,
it is potentially more sensitive than ELISA for detecting leafroll
virus from current season infection.
This study was performed to determine 1) how quickly PLRV moves
to the tubers when plants are infected at different ages and times
during the season and 2) how reliably DTBA detects PLRV in primary
infected vines and tubers.
34
Methods and Materials
Field plot layout and seed preparation
Russet Burbank seed potatoes were planted in a randomized
complete block design and sampled according to a split-block
arrangement at the Lewis-Brown Horticultural Farm near Corvallis,
This site in the Willamette Valley was chosen despite high
Oregon.
aphid populations because of the relative absence of local potatoes and
PLRV inoculum.
Plots were replicated six times in 1991 and three in
1992.
Certified, generation three seed was planted both years.
Certification readings for PLRV and mosaic viruses in the field and in
winter grow-out tests were zero for both seed lots.
Seed was cut on
May 6, 1991 but planting was delayed until May 24 because of heavy
rains.
Seed for 1992 trials was cut three days before planting on May
22 and June 8.
Cut seed was allowed to suberize at room temperature in
Seed pieces
1992 but was held in 10 °C storage until planting in 1991.
averaged 1.5 to 2.0 ounces both years.
Crop production practices
Fertilizer (15-15-15) was broadcast at 560 kg/ha preplant and
banded at 481 kg/ha on both sides of the furrow at planting.
row was planted with 30 seed pieces spaced 23 cm apart.
cm apart.
Each plot
Rows were 86
Each plot consisted of two planted rows bordered by two
blank rows and two outside border rows.
Border rows were treated with
aldicarb insecticide at 10.9 kg/ha before plant emergence.
meter bare ground alley was left between blocks.
A three
A border row was
planted through the middle of each alley and treated with aldicarb.
This arrangement provided bare ground and insecticide treated barriers
around each plot to hinder movement of apterous (non-winged) aphids.
Weeds were controlled by one application of metribuzin (0.6 kg/ha) at
35
early plant emergence both years.
sprinklers as needed.
Plants were irrigated with solid set
A yellow pan water trap was placed in each
corner of the field to monitor alate (winged) aphid movement.
were collected and fresh water was added to pans weekly.
Aphids
Fungicide
sprays of chlorothanil (Bravo) were alternated with metalxyl (Ridomil)
as needed for control of early and late blight during August.
Inoculation of field-grown plants with viruliferous aphids
Apterous Myzus persicae were generously supplied by Dr. Harold
Toba and Mr. Lee Fox of the USDA-ARS Entomology Research Laboratory in
Yakima, Washington.
This aphid colony was initiated March 15, 1985
from a fundatrix (stem-mother aphid) obtained from a peach tree.
The
colony has since been maintained on Physallis floridana or Datura
The leafroll virus strain
stramonium plants in the lab or greenhouse.
is PLRV 2243, isolated from cv. Russet Burbank plants in the Yakima
area (Fox et al., 1993).
Aphids reared on PLRV-infected plants were
shipped to Oregon State University on live plants or detached leaves
and placed in the field within one week of their arrival in Corvallis.
Winged aphids were destroyed prior to transferring non-winged aphids to
Before aphids were placed
plants with sable-bristled artists brushes.
on plants, stems were thinned to one per hill so that all tubers were
equally liable to infection.
Ten to twenty aphids were placed on each
plant at upper, middle, and lower leaf axils.
The inoculation access
period was limited to two days in 1991 and three in 1992.
The entire
planting was sprayed with methamidophos or oxydemeton-methyl at the end
of each inoculation period.
The inoculated plots were then sidedressed
with aldicarb and all plots were watered.
Aldicarb was applied to
inoculated plots to minimize the spread of PLRV to non-target plots.
Aphids require about an hour to acquire PLRV from an infected
plant (Harrison, 1984; Kostiw, 1991) and 26% of the aphids exposed to
aldicarb treated potato plants are killed after one exposure (84% after
36
four exposures) (Holbrook, 1977).
It would therefore be possible for
an aphid to acquire the virus but the probability of infecting plants
in other plots under the circumstances employed was minimal.
Plots were kept relatively free of aphids by spraying every other
week with methamidophos in 1991 and a rotation of oxydemeton-methyl
(Metasystox-R), methomyl (Lannate), and methamidophos (Monitor) in
1992.
Pyrenone (6% pyrethrin and 60% piperonyl butoxide) was added at
two ounces per acre to many of the 1992 sprays because of a desirable
fuming action which forces insects out of sheltered areas and into
contact with spray residue.
The very cold winter temperatures of
1990-91 substantially reduced spring aphid populations in 1991 compared
to the levels of 1992 (Figure 3.3).
at emergence in 1992.
Aphids were found on most plants
Insecticide sprays were applied whenever a
random examination of plants in the field revealed viviparous females
deposited by winged aphids.
Plants were inoculated on June 6 at 33 days after planting (DAP)
and July 11 at 48 DAP in 1991.
In 1992, plants were inoculated on July
4 at 42 DAP, on July 22 when two different lots of plants were 44 and
61 DAP, and August 10 at 63 DAP.
Plant growth stages corresponding to
inoculation date are shown in Table 3.7.
Results from 1991 showed a sharp decline in infection levels
between the first and second inoculation dates.
Therefore, treatments
were expanded in 1992 to determine whether plant age (DAP) or
inoculation date most strongly influenced plant infection levels.
First inoculations were performed when skies were overcast and
temperatures were low both years.
high temperatures and clear skies.
Later inoculations coincided with
Aphids were applied in the early to
late afternoon during the later inoculations to facilitate rapid
acclimation and feeding during the cool night hours.
37
Sampling of plants
Plant samples were collected at 5, 9, 13, 17, and 21 days after
In 1991, 4
inoculation and on two additional dates in July, 1991.
plants were sampled from each plot resulting in 24 plants from 6
replications; 10 plants were collected from 3 replications per date for
a total of 30 samples in 1992.
Plants were taken to a field lab where
stems were sectioned with a fresh razor blade into upper, middle, and
basal portions.
Cut stem ends were blotted onto nitrocellulose
membranes (NCM) for DTBA tests for PLRV (Chapter 2).
weighed and assigned identification numbers.
Tubers were
Approximately one cm of
the stolon end of each tuber was removed and sliced longitudinally
through the stolon attachment.
then blotted onto the NCM.
Tissues exposed by the last cut were
The tuber cutting pattern permitted
detection of PLRV at the stolon end, the PLRV entry point.
Tubers from
each plant were placed in a sleeve of yellow mesh netting and separated
by twist-ties so that individual tuber identity could be maintained in
storage and winter grow-out tests.
Samples were blotted on NCM and
processed according to DTBA methods listed in Chapter 2.
Grow-out tests for PLRV confirmation
Tuber grow-out tests were performed during the winter to confirm
summer results.
Tubers were stored at 4.4 °C in year one and at 10 °C
in year two to promote dormancy break and avoid possible freezing
injury to small tubers.
Tubers were stored for three months in 1991
and two months in 1992 before dormancy breaking treatments.
Tubers
were dipped in ammonium thiocyanate (20 g/l) for one hour and held in a
warming room at 26 °C for two weeks to help break dormancy.
They were
then indexed by excising tuber bud ends with a melon-ball scoop and
inserting a small numbered metal tab into each.
The metal tab number
corresponded to the tuber identification number assigned at harvest.
The melonball seed pieces were sprayed with a five percent chlorox
38
solution and allowed to suberize overnight at room temperature before
planting in soil beds.
When plants were at least 20 cm tall, visual
PLRV readings were recorded and plant stems were sampled at upper and
lower regions for DTBA testing.
Visual symptoms were evaluated by the
author and O. Gutbrod, Oregon State University Seed Potato
Certification Specialist.
Plants representing tubers from the first
inoculation of each year were also tested with ELISA for PLRV.
Tissues
used for DTBA were also used for ELISA to facilitate comparison of the
DTBA and ELISA results.
Inoculation and detection of PLRV in tissue culture-derived plantlets
Two sets of cv. Russet Burbank clones were started from diseasetested tissue cultured plants grown under a 16 hour photoperiod (187
pmol
m-2)
in the laboratory.
The first set was planted Dec. 17, 1992
and inoculated on Jan. 7, 1993.
The second set was planted Jan. 26,
and inoculated on Feb. 3, 1993.
Plants were grown in screen cages in a
greenhouse and had 16 hours of supplemental light (95 pmol s' m-2) per
day.
Plants were inoculated in the laboratory.
Aphids and the PLRV
source were obtained from Dr. Harold Toba and Lee Fox in March, 1991
and maintained on infected Physallis floridana plants until used for
the 1993 inoculations.
Three to five aphids were placed on the top
leaves of each plant and were confined by inverting a modified 15 dram
clear plastic vial over the top and placing the cap, with a hole cut
for the stem, on the vial.
The stem hole in the vial cap was plugged
with cotton to prevent aphid escapes.
bamboo stake for support.
The dram was then fastened to a
After six days, aphids were counted and then
killed by placing plants in a fumigation chamber and using Resmithrin
(1.0% resmithrin active ingredient) or X-clude, an encapsulated natural
pyrethrum (both products from Whitmire Research Laboratory, St. Louis,
MO).
After inoculation, plants were placed under screen cages in the
greenhouse.
At 7, 10, 13, 16, and 19 days after inoculation, each stem
39
region containing an axillary bud was sectioned and tested for PLRV
with the DTBA.
Each stem cutting was numbered and placed in a 72-count
flat of aggregate soil mix (Sunshine-mix #4; Fisons, Vancouver, B.C.,
Canada) for a grow-out test.
When plantlets from the stem cuttings
were at least 24 days old they were tested for PLRV with the DTBA.
At
all times plants were kept under screen cages and the greenhouse was
monitored for aphids.
Statistical analyses
Results of winter grow-out and summer tests were statistically
compared using a chi-square test for paired samples.
Test data were
discrete since plants were either PLRV positive or negative.
The chi-
square test for paired samples was described by the equation X2= (la-bl1)2/n where n=a+b.
Letters "a" and "b" represent the number of tubers
which tested positive for DTBA and negative for ELISA, or vice versa
(Snedecor and Cochran, 1977a).
This test allowed for statistical
comparison of the tubers that produced different results for each
assay.
Tuber weights from PLRV-infected plants were compared by use of a
t-test which accounts for unequal sample size and variance as described
by Snedecor and Cochran (1977b).
40
Results
Plant age and inoculation date effects
Winter grow-outs of tubers harvested at different times during
the growing season show that the average number of days required for
PLRV to move from inoculation sites to the tubers increased with both
plant age and delayed inoculation.
PLRV was detected in tubers by DTBA
about three days after detection in vines for both 1991 inoculations
(Table 3.1).
Inoculations 17 days apart in 1991 showed a marked decrease in
plant and tuber infection between 33 and 48 days after planting.
Winter grow-out tests of tubers suggested that almost every tuber in a
hill became infected with early inoculations (June 26, 33 DAP) while
only 50% (at 21 days post-inoculation) were infected in plants
inoculated 17 days later (Table 3.2).
Plant and tuber infection levels also declined when inoculations
were delayed by 18 to 20 days in 1992.
As in the previous year, the
earliest 1992 inoculation produced the highest percentage of infected
plants and tubers (Table 3.2).
PLRV inoculation was less effective and
titre levels developed more slowly in 1992 than in 1991 as evidenced by
delayed detection and lower infection levels.
When plants of different
ages were inoculated on the same date (22-Jul-92), infection levels
were lowest among older plants (Table 3.2, Figure 3.2).
Similar
declines in infection levels with age were observed when inoculation
was delayed by 18 to 19 days; the later inoculation date produced fewer
infected plants than did the earlier date.
Reliability of DTBA for detecting PLRV translocation in field-grown
plants
Plant foliage from winter grow-outs of tubers give the most
reliable ELISA results for PLRV (Flanders et al., 1990); reliability is
probably enhanced because small, undetectable levels of virus present
41
in the tuber at harvest can replicate to a detectable level during
sprouting and emergence.
Comparison of results of winter grow-out tests with DTBA, ELISA,
and visual inspection readings showed the summer DTBA test of tuber
tissue to be approximately equal to winter DTBA stem tests in the first
year.
A winter DTBA test detected two more PLRV positive tubers from
the early (33 DAP; 26-Jun-91) inoculation than did a summer DTBA and
one more than a winter ELISA, an insignificant difference (Table 3.3).
Winter ELISA and DTBA tests were all done with stem tissue and
results were virtually identical for the two methods.
This finding
confirms results described in chapter two, which show ELISA and DTBA to
be equally precise for secondary PLRV detection in stem tissue.
Summer
DTBA tests of tuber tissue always detected fewer positives than winter
grow-out tests of stems from those same tubers.
DTBA results from field grown, PLRV-inoculated plants were
distinct and easy to interpret (Figure 3.4).
Purple color reactions
confirm that PLRV is confined to the phloem in stem and tuber tissues.
Reliability of DTBA for detecting PLRV in tissue culture derived
plantlets
Potato leafroll virus was detected with DTBA when plant stems
were sampled and again when axillary buds from those stem sections were
grown out into plantlets.
The stem section tests represent primary
PLRV as it enters the plant.
The planlets from axillary buds also
represent primary PLRV, but provide a second test of the same tissue
section.
The second test of the plantlet should allow more accurate
detection of PLRV as the virus would have more of a chance to replicate
to a detectable level.
At seven days post inoculation more PLRV was
detected in the first test than in the second test and the reverse is
true at 19 days post inoculation (Table 3.4).
42
PLRV effects on yield
Tuber yields from infected plots harvested in late September were
usually lower than those from the healthy control plots (Table 3.5).
In 1991, the relationship (r2) between yield per plant and plant
infection percentages was moderate, while tuber size and number per
plant were strongly related to the percentage of PLRV-infected plants
(Table 3.6).
Similar trends were observed in 1992, but less strongly,
especially between yield per plant and the incidence of PLRV
infections.
tubers.
The healthy control plots always produced fewer but larger
Plots inoculated early in the season produced the highest PLRV
levels and the largest percentage of small tubers.
Not all of the tubers from an infected plant contained PLRV, at
least at detectable levels.
healthy tubers.
Infected tubers were always larger than
However, the difference in average tuber size between
infected and healthy tubers was significant (p=0.05) for only one set
of samples 21 days post-inoculation in 1992 (Table 3.7).
The fact that
larger tubers probably formed first and therefore were exposed to PLRV
longer may have accounted in part for higher infection percentages.
The use of aldicarb-treated guard rows and frequent insecticide
sprays prevented the spread of PLRV from infected plots.
examination showed little PLRV spread among plots.
Careful
Some spread into
the control plots did occur in the early 1992 planting (Table 3.5), but
other control plots contained no detectable PLRV.
43
Discussion
The average time lapse between inoculation and tuber infection
ranged from 11.7 days to more than 21 days depending on inoculation
date and plant growth stage.
Tuber infection occurred within nine days
of inoculation in one instance.
These results suggest that PLRV is
translocated to the tubers more slowly and less efficiently with late
season inoculation of mature plants (Table 3.2).
The exception in this
study was that more plants became infected in older plots (-62 days
after planting) when they were inoculated late in the season compared
to an earlier inoculation (Figure 3.2).
Storch and Manzer (1985)
reported that PLRV introduced late in the season reached the tubers
within three days (5.2% of plants had infected tubers) and that 30.3%
of the plants had infected tubers by 15 days after inoculation.
A
possible explanation for the apparent time discrepancy between these
two studies may involve the duration of aphid feeding.
In this study,
aphid access to the plants was limited to two or three days while
Storch and Manzer did not mention aphid removal or insecticide
applications.
An unlimited aphid feeding period would produce much
higher virus pressure.
When Russet Burbank potatoes are planted early in the season and
PLRV and vectors are present, plants are very susceptible to PLRV
infection.
In plants infected at 33 DAP, tuber infection approached
100% within 21 days.
Plants inoculated 15 days later were less
susceptible to infection and fewer tubers became infected (Table 3.2;
Figure 3.1).
It is possible that climatic factors altered aphid
behavior and infection levels between the two dates since aphid
behavior is weather-dependant.
However, environmental influences can
be discounted in comparisons of different age plants inoculated on the
same day (Table 3.2; see 44 and 61 DAP).
In the same-date inoculation,
younger plants were more susceptible than older ones (planted 17 days
earlier) confirming plant age effects on infection levels.
44
Infection trends for similar age plants inoculated at different
chronological dates are less clear.
In these inoculations, age can be
discounted because plants were not only similar in age, but also in
growth stage (Table 3.8).
In young plots the early inoculation (-43
DAP) produced the highest percentage of PLRV-infected plants.
Conversely, late inoculation (-62 DAP) produced the highest percentage
of PLRV plants in older plots (Figure 3.2).
The older plots had a
larger average tuber size at inoculation, suggesting a different stage
of tuber bulking, which might account for higher PLRV levels compared
to the early inoculation.
Other studies have generally shown that late inoculations produce
low infection rates (Cadman and Chambers, 1960; Flanders et al., 1990;
Knutson and Bishop, 1964).
Storch and Manzer's (1985) study included
both plant age and inoculation date as factors.
Their results suggest
that neither contributes significantly to the level of PLRV in cv.
Russet Burbank plants but both significantly effect PLRV levels in cv.
Katandin.
However, all of Storch and Manzer's inoculations occurred
within a one month period from August 20 to Sept. 16 and are therefore
essentially all late season inoculations.
In this study fewer mature plants became infected than younger
plants and the number of infected tubers on more mature plants was
greatly reduced.
Most seed growing areas are isolated, have short
seasons, and are subject to mid-to-late season in-flights of aphids
corresponding to the mid-to-late season inoculations in this and
related studies.
In a seed growing area, better protection against
PLRV could probably be achieved by planting as early as possible so
that plants are older when the aphid flights typically begin.
Current season (primary PLRV) was detected with DTBA as early as
seven days post inoculation in tissue culture derived greenhouse-grown
plantlets.
A test of plant tissue grown from infected tissue (source
tissue) should be a more accurate test of PLRV infection.
Even small
45
undetectable amounts of PLRV have a greater likelihood of replicating
to detectable levels if the plant source tissue is infected because the
infection is continous.
Results from these tissue culture plants are
inconclusive, but they do show that detection of primary PLRV with DTBA
is not consistent (Table 3.4)
Summer DTBA tuber (stolon end) results at 21 days postinoculation were not significantly different than results from winter
grow-out tests of the same tubers.
This is true for all inoculation
dates except 61 DAP (22-Jul-92, Table 3.3) which produced more summer
positive tests than winter positive tests, suggesting that some summer
tuber tests may have produced false positives.
There is some
difficulty involved in the interpretation of DTBA tests of tuber tissue
(Table 2.1) and more reliable readings are provided by stems or
petioles.
Even though statistically significant differences were not
found between summer and winter DTBA tests of tubers harvested 21 days
post-inoculation, most summer DTBA detected fewer positive tubers than
did winter DTBA (Table 3.3).
A DTBA test could be effectively used on
a sample of tubers at the end of the season to estimate the incidence
of PLRV in the crop, but a winter grow-out test using the stem tissue
for DTBA (chap. 2), or leaf tissue (Flanders et a/, 1990) for ELISA
A
still remains the most accurate measure of PLRV infection levels.
strong correlation between visual symptoms and DTBA readings in the
winter tests was noted thus providing a second independent confirmation
of DTBA results.
Knutson and Bishop (1964) reported an 18 % greater PLRV infection
frequency among large tubers in a hill than among small tubers.
They
recommended using tubers larger than 57 g to 113 g (2-4 oz.) in winter
grow-out tests to avoid underestimating PLRV levels.
Results from the
present study showed a similar association between tuber size and PLRV
infection in that healthy tubers always had a smaller average size than
infected tubers (Table 3.7).
However, no recommendation can be made
46
about optimum tuber size for winter tests because data were based on
tubers harvested after inoculation but before typical commercial
harvest.
47
Summary and Conclusions
Planting date, as related to Russet Burbank plant age and growth
stage at PLRV infection, has a more consistent influence than the
actual inoculation date on infection levels.
Late season feeding by
viruliferous aphids transmits PLRV but to fewer plants than early
season feedings, similar to results obtained in "mature plant
resistance" studies.
More emphasis should be placed on protecting
plants when aphids are present early in the season and plants are most
susceptible.
The early use of insecticides and/or row covers would be
especially effective in controlling virus spread.
DTBA of stems and petioles is a useful, reliable tool for
detecting tuber-borne (secondary) PLRV but less useful for detecting
current season (primary) PLRV.
ELISA, DTBA, and visual inspection are
comparable for detecting PLRV in winter grow-out tests commonly used by
Certification Programs.
ELISA or DTBA can be used to confirm
questionable visual symptoms.
Either serological assay could be used
to confirm visual evaluations when training new certification
inspectors or to
confirm PLRV symptoms in new varieties.
Although
DTBA does not quantify virus levels as does ELISA, it accurately
indicates whether or not a plant is infected and is easier to use than
ELISA for that purpose.
DTBA can detect current season (primary) PLRV in field-grown
potato vines and tubers as early as nine days after infection.
Primary
PLRV can be detected with DTBA in tissue culture derived greenhousegrown plantlets as early as seven days after infection.
primary PLRV less reliably than secondary PLRV.
DTBA detects
DTBA is as reliable as
ELISA for tuber-borne (secondary) PLRV in stem tissue.
Potato stems
could be tested near the end of the season to estimate PLRV levels and
predict results of winter grow-out tests.
If a grower suspects a seed
lot is near the tolerances allowed for recertification, a DTBA estimate
of PLRV percentages would help in deciding whether to submit the lot
48
for winter certification tests or sell it for production of a
commercial (ware) crop.
More research on late season DTBA testing of
plant and tuber tissue for PVX and PVY is needed to evaluate its
usefulness for these viruses.
When young plants become infected early in the season, nearly all
of the tubers in a hill will become infected.
Larger tubers in the
hill are more prone to infection than small tubers.
49
Table 3.1. Average number of days post-inoculation required for
detectable potato leafroll virus (PLRV) infections in stems and
tubers.
PLRV
actually
Days to first detection
of PLRV by DTBAI
present2
(days)
Inoculation
date
Plant age
(days from
planting)
26-Jun-91
33
11.0
14.3
14.3
11-Jul-91
48
22.5
25.5
22.3
4-Jul-92
42
14.3
14.3
11.7
22-Jul-92
44
19.7
19.7
13.0
22-Jul-92
61
nd3
nd
>21.0
10-Aug-92
63
nd
nd
>21.0
Stems
Tubers
Tubers
1 represents current season primary infection
2 represents secondary infection; from winter grow-out tests
3 nd - not detected in all replications by 21 days after planting
Percentage of Russet Burbank plants and tubers infected with potato leafroll virus as
Table 3.2.
determined by the Direct Tissue Blotting Assay.
Number of
Plant age
% Infected tubers2
tubers
% Infected plants'
Number of
(days
from
Days postplants
from
Inoculation
Winter
Summer
Winter
Summer
infected
sampled
planting) inoculation
date
plants
0
0.0
0.0
0.0
0.0
24
5
33
26-Jun-91
0.0
100.0
8
12.5
29.2
24
9
96.7
30
70.8
70.8
73.3
24
13
88.9
95.6
45
58.3
58.3
24
17
93.2
88
90.9
95.8
95.8
24
21
11 -Jul -91
48
5
9
13
17
21
28
35
4-Jul-92
42
5
9
13
17
21
23
24
24
24
24
24
24
0.0
0.0
0.0
12.5
8.3
29.2
20.8
0.0
4.2
0.0
12.5
8.3
29.2
20.8
30
30
30
30
30
0.0
0.0
16.7
23.3
20.0
0.0
3.3
40.0
26.7
26.7
0
2
0
10
12
11
18
0
1
50
36
39
0.0
0.0
0.0
40.0
25.0
63.6
55.6
0.0
50.0
0.0
60.0
50.0
72.7
88.9
0.0
0.0
14.0
55.6
53.6
0.0
na 3
68.0
83.3
69.2
Table 3.2. Continued
Plant age
Inoculation
date
22 -Jul -92
(days
from
planting)
44
Days postinoculation
5
9
13
17
21
22 -Jul -92
61
5
9
13
17
21
10 -Aug -92
63
5
9
13
17
21
1
2
3
Number of
plants
sampled
%
Infected plants'
Summer
Winter
30
30
30
30
30
0.0
3.3
0.0
6.7
16.7
0.0
6.7
3.3
6.7
20.0
30
30
30
30
30
0.0
3.3
0.0
0.0
3.3
0.0
3.3
0.0
0.0
6.7
30
30
30
30
30
0.0
0.0
13.3
0.0
0.0
13.3
6.7
10.0
6.7
6.7
Number of
tubers
from
infected
plants
0
4
4
10
17
0
7
0
5
11
0
0
16
12
20
% Infected tubers2
Summer
Winter
0.0
0.0
0.0
40.0
52.9
0.0
25.0
25.0
40.0
64.7
0.0
14.3
0.0
0.0
0.0
0.0
0.0
0.0
0.0
9.1
0.0
0.0
12.5
0.0
5.0
0.0
0.0
0.0
0.0
15.0
Plants or tubers scored positive; summer = stems or tubers positive; winter = stems grown from
tubers
Summer = tuber tissue (stolon end); winter = stem tissue grown from summer-tested
tubers
tubers decayed
infected Russet Burbank tubers' as determined by
Table 3.3. Number of potato leafroll virus (PLRV)
Immunosorbent
Assay (ELISA).
Tissue Blotting Assay (DTBA) and Enzyme-Linked
Winter DTBA
Summer DTBA vs.
Number of PLRV infected
vs. winter:
winter:
tubers
Winter
Plant age
No. of
Days Summer
(days
InoculVisual ELISA
DTBA Visual ELISA
post
from
DTBA
ation
DTBA Visual ELISA tubers
sampled2
inoc.
planting)
date
26-Jun-91
11-Jul-91
4-Jul-92
33
48
42
5
9
0
0
ERR
ERR
*
*
*
NS°
NS
NS
NS
NS
NS
NS
NS
NS
NS
NS
NS
NS
NS
NS
NS
NS
84
84
63
70
88
69
92
ERR
ERR
NS
NS
NS
NS
NS
NS
NS
0
6
ERR
ERR
1
NS
NS
ERR
ERR
ERR
NS
ERR
ERR
ERR
35
107
**
**
**
8
28
43
81
0
0
0
0
1
1
13
17
21
28
35
0
0
4
6
2
6
3
6
7
7
6
8
10
15
12
5
9
0
0
0
0
0
17
29
29
20
21
ERR
**
8
5
9
7
ERR
*
9
29
43
82
13
17
21
ERR
*3
0
22
40
80
1
34
30
27
vs.
winter
ELISA
ERRS
0
13
17
21
Winter
visual
0
20
42
76
89
0
27
43
80
Direct
30
32
28
160
159
ERR
NS
NS
NS
*
ERR
ERR
ERR
NS
ERR
ERR
ERR
NS
*
*
**
**
NS
*
*
NS
NS
NS
NS
NS
NS
**
NS
NS
Table 3.3.
Continued
Plant age
Inoculation
date
from
planting)
22 -Jul -92
44
22 -Jul -92
(days
61
Number of PLRV infected
tubers
Winter
Days Summer
Post DTBA
DTBA Visual ELISA
inoc.
5
0
0
0
9
0
0
4
9
1
1
1
13
17
21
4
3
11
10
5
0
2
2
0
0
0
0
0
9
13
17
21
10-Aug-92
63
5
9
13
17
21
0
9
1
0
0
0
0
0
0
0
O
0
0
0
0
0
2
1
3
2
0
0
2
Summer DTBA vs.
winter:
No. of
tubers
sampled2
47
94
115
107
112
Winter DTBA
vs. winter:
vs.
DTBA
Visual
ERR
ERR
ELISA
Visual ELISA winter
ELISA
NS
NS
NS
NS
NS
ERR
NS
NS
266
244
178
165
205
ERR
ERR
NS
NS
NS
NS
NS
NS
NS
118
105
115
105
147
ERR
ERR
NS
ERR
*
NS
ERR
ERR
NS
NS
NS
ERR
ERR
ERR
ERR
NS
ERR
ERR
ERR
ERR
NS
NS
NS
NS
NS
NS
1 Summer = tuber tissue (stolon end); Winter = stem tissue grown from summer-tested tubers
2 Includes tubers from all plants (healthy and infected)
Chi-square test for paired samples
3 * p=0.05, ** p=0.01 (significant at these levels)
4 NS not significant
5 ERR all tubers had the same results for compared tests
Winter
visual
54
Table 3.4.
Potato leafroll virus (PLRV) infection percentages
in stems and nodal cuttingsl of Russet Burbank plants inoculated
by viruliferous Myzus persicae.
Number of PLRVpositive tests
Days postinoculation
Stem
Plantlet
Percentage of
PLRV positive
stem and
plantlet tests
that match
Number
of
samples
7
12
8
17.7
17
10
1
9
0.0
10
13
10
8
63.6
11
16
8
14
40.0
15
19
27
31
81.1
32
1 Each stem section with an axillary bud was tested post-inoculation,
rooted, and the resulting plantlet was tested at least 24 days later.
Data represent two separate experiments.
55
Yield components from fall-harvested plants with different
Table 3.5.
levels of potato leafroll virus (PLRV) and from different planting
dates.
Plant age at
Tuber
Average
Average
inoculation
yield/
number of
tuber
% Plants
(days from
tubers/plant plant (g)
size (g)
with PLRV
planting)
Year
1991
33
48
control
1992
(early
planting)
control
142.5a'
56.7
21.5
9.1
232.7a
329.7ab
341.2 b
218.9b
226.9c
6.4a
5.2b
5.1b
13.6
10.1
8.9
909.3a
1081.2ab
1139.7 b
2955.6
2887.9
3040.6
2098.5
7.3
277.6
1698.0
7.1
238.3
1820.2
6.1
321.4
control
within
each
block
are
not
Numbers followed by same letters
significantly different at p=0.05 level. Numbers not followed by
letters within each block are not significantly different.
1992
(late
planting)
1
42
61
73.6
20.4
0.0
44
63
36.7
26.5
0.0
Correlation coefficients (r2) for assessing relationships
Table 3.6.
between percentages of plants with potato leafroll virus (PLRV) at
harvest and yield components.
% PLRV Plants
1992 (late
1992 (early
1991
planting)
planting)
-0.47
-0.45
-0.81
Tuber size
0.53
0.65
0.81
Number of tubers per
plant
-0.01
-0.20
-0.61
Yield per plant
56
Table 3.7.
Average tuber size (g) for infected and healthy tubers
from Russet Burbank plants testing positive for potato leafroll virus
(PLRV) using the direct tissue blotting assay.
Inoculation Plant age
Days
Average tuber weight (g)
date
(days from
post
PLRV Healthy
planting) inoculation
infected
11-Jul-91
48
19
42.8
28.8
NS
21
75.5
36.1
NS
28
158.5
89.4
NS
35
182.9
115.0
NS
4-Jul-92
42
17
19
21
24.1
45.2
83.0
17.5
37.6
41.7
NS
NS
*
61
22-Jul-92
61
143.5
99.0
NS
Significant at p=0.05 using t test based on Snedecor and Cochran
(1977b) method for unequal variances and sample sizes
NS not significant
*
57
Table 3.8.
Russet Burbank plant growth parameters at potato leafroll
virus inoculation by viruliferous aphids.
Inoculation
date
Plant age
(days from
planting)
Vine
length
(cm)
Nodes
per
plant
Tubers
per
plant
Average
tuber
weight (g)
26-Jun-91
33
15.0
8.4
0.3
na
11- Jul -91
48
46.2
14.1
3.9
13.8
4-Jul-92
42
47.2
14.1
0.1
na
22-Jul-92
44
46.2
16.7
0.9
6.3
61
77.2
21.1
5.2
73.0
63
86.9
23.8
4.0
115.2
10-Aug-92
58
100
90
80
70
oG
O
60
z
50
a
40
114
30
20
10
0
5
13
9
21
17
DAYS FROM INOCULATION TO HARVEST
SUMMER FIELD TEST
WINTER GROWOUT
100
90
80
[48
70
DAP; 11 Jul -91
60
50
40
30
20
10
oolo
0
5
9
13
17
21
28
35
DAYS FROM INOCULATION TO HARVEST
SUMMER FIELD TEST
WINTER GROW-OUT
Figure 3.1. Percentage of potato leafroll virus positive plants
inoculated at 33 and 48 days after planting (DAP).
Summer test results
for stems and tubers; winter test results for stems of plants grown
from summer-tested tubers.
59
40
35
30
42 DAP; 4-Jul-92
44 DAP; 22-Jul-92
25
20
N
15
10-
04
5
9
13
17
21
5
9
13
17
21
DAYS FROM INOCULATION TO HARVEST
40 -*
35
30 -
61 DAP;
22-Jul-92
63 DAP; 10-Aug-92
25
20
15
10
9
13
17
21
13
17
21
DAYS FROM INOCULATION TO HARVEST
SUMMER FIELD TEST 1111 WINTER GROW-OUT
virus infection of plants
Figure 3.2. Percentage potato leafroll
inoculated at similar days after planting (DAP) (similar growth
stages), but on different calendar dates. Summer test results based on
stems and tubers; winter test results based on stems of plants grownout from summer-tested tubers.
60
350,-
300
1991
250
200
150
100
50
0
,L
'
71-----7--11-111--0
20-Jul
6-Jul
19-Jun
12-Jul
29-Jun
II
7-Sep
23-Aug
10-Aug
31-Aug
16-Aug
2-Aug
350
300
250
200
150
100
50
1
1
18-Aug 31-Aug
3-Aug
21-Jul
6-Jul
22-Jun
8-Jun
8-Sep
11-Aug
25-Aug
27-Jul
14-Jul
29-Jun
15-Jun
-41F- GREEN PEACH APHIDS --6E- OTHER APHIDS
Figure 3.3. Natural aphid populations in Russet Burbank plots at
Corvallis, Oregon. Aphids were collected weekly from water-filled
yellow pan traps located in each corner of the field.
61
Processed nitrocellulose membranes showing typical PLRV
Figure 3.4.
reactions from inoculation on June 26, 1991 (33 days after planting).
Stem and tuber blots from (A) plant samples 5 days post-inoculation;
Controls at the bottom of
(B) plant samples 17 days post-inoculation.
each membrane are 'A', 'B' FLRV-infected; 'C', 'D' healthy.
62
Literature cited
Cadman, C.H. and J. Chambers. 1960. Factors affecting the spread of
aphid-borne viruses in potato in Eastern Scotland. Ann. Appl.
Biol. 48:729-738.
Flanders, K.L., D.W. Ragsdale, and E.B. Radcliffe. 1990. Use of enzymelinked immunosorbent assay to detect potato leafroll virus in
field grown potato, cv. Russet Burbank. Amer. Potato J. 67:589602.
1985.
Francki, R.I.B.
Florida.
Atlas of plant viruses.
CRC Press, Coca Raton,
Potato leafroll virus - Effect of
Knutson, K.W. and G.W. Bishop. 1964.
date of inoculation on percent infection and symptom expression.
Amer. Potato J. 41:227-238.
Effect of time and date of
1985.
Storch, R.H. and F.E. Manzer.
inoculation, plant age, and temperature on translocation of
Am Potato J 62:137potato leafroll virus into potato tubers.
43.
63
General Conclusions
Potato leafroll virus (PLRV) causes yield and quality losses in
Russet Burbank and other susceptible potato cultivars.
Russet Burbank
accounts for a majority of the processed acreage in the Pacific
Net necrosis, characterized by necrotic tuber tissue, is
Northwest.
strongly associated with PLRV infections and a common quality defect in
Russet Burbank.
Enzyme-linked immunosorbent assay (ELISA) has been used for
about ten years by many certification programs to test potato plants
for viruses.
The direct tissue blotting assay (DTBA), based on the
same principles as ELISA but using undiluted plant sap blotted onto
nitrocellulose membranes, offers a quick and simple alternative to
ELISA.
DTBA, as tested in these studies, is as accurate and reliable
as ELISA for detecting tuber borne (secondary) PLRV as long as stem or
petiole tissues are used.
DTBA is not as reliable as ELISA for testing
leaflets and while it cannot quantify virus titre it is effective for
detection of PLRV. Detection alone is sufficient for certification
purposes.
Control of PLRV in seed fields is obtained by the use of healthy
seed, removing diseased plants, and controlling aphid vectors with
insecticides.
Seed potato growing areas are usually isolated and not
favorable to overwintering aphids.
Consequently, aphid flights usually
occur later in the season than in commercial productin areas.
When
inflights of aphids begin late in the season, some areas recommend or
require removal or killing of potato vines to stop movement of PLRV to
tubers.
The time lapse between vine and tuber infection is reported to
vary from 3 to 10 days.
Differences in virus sources, plant age, and
duration of aphid feeding can affect the rate of PLRV movement.
In
64
these studies, PLRV was detected by DTBA as early as nine days after
inoculation.
Plants inoculated early in the season and/or at a young
age were more susceptible to PLRV infection than plants inoculated
later and/or at an older age.
Age more consistently effected PLRV
levels in a plot than did the actual calendar date of inoculation.
Russet Burbank plants developed a considerable degree of resistance to
This result suggests that in areas where
infection as they matured.
inflights of aphids occur late in the season, early planting would
reduce infection levels since plants would be more mature when
subjected to PLRV inoculation.
Regardless of when aphids normally
appear, careful attention should be paid to the use of healthy seed,
removal of diseased plants, and aphid control.
Further studies should focus on making DTBA more useable by
growers to allow better on-farm estimates of PLRV levels and other
viruses in their seedlots.
A seed grower using DTBA to obtain an
estimate of PLRV levels might decide it is cost effective to forgo the
expense of winter certification testing if a seed lot is at or near
tolerance levels toward the end of the season.
When Russet Burbank is
infected with PLRV, net necrosis is likely to develop in storage making
the crop less valuable.
A decision to market an infected seed crop to
a processor before net necrosis develops or before the lot is rejected
in the winter certification test would save the grower money.
Certification programs should altho evaluate DTBA as a replacement for
ELISA for large scale testing of seed lots.
While DTBA has been shown
to be effective for detecting PVX, PVY, and PLRV (Samson et a/, 1993;
also see chapter 2 in this thesis), research is needed to quantify
virus titre with DTBA.
65
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