- Email: [email protected]
Genetic control of 32P transfer from wheat to Erysiphe graminis f. sp. tritici during primary infection
PhysiologicalPlant Pathology(1978) 13, 1-l 1
Genetic control of 32Ptransfer from wheat to Erysiphe gram inis f. sp. frifici during primary infection T. J. MARTIN? and A. H. ELLINGBOE Departmentof Botany and Plant Pathology, Michigan State University East Lansing, Michigan 48824, U.S.A. (AcceptedforpublicationJanuary 1978)
Environmental conditions necessary for synchronous development of Etysiphe graminis D.C. f. sp. tritici Em. Marchal during primary infection of wheat were shown to affect the amount of szP taken up and translocated to the epidermis of excised leaves placed in a saP solution. Light increased transpiration, the amount of 32P taken up by the leaves and the amount of s2P transferred to the parasite. Rates of saP transfer from host to parasite were determined for compatible and incompatible parasite-host interactions during primary infection. Transfer rate reflected the relative compatibility of the host-parasite interaction studied. Interactions with little fungal development had lower total rates of transfer than did compatible interactions. However, when transfer was expressed as saP transferred per successful primary infection, the successful parasite units involved in incompatible interactions obtained saP at a greater rate than did parasite units in compatible interactions.
INTRODUCTION
The primary infection process of Erysiphe graminis on wheat (T&cum aestivum) has been divided into distinct morphological stages : (1) germination, (2) production of club-shaped appressorial initials, (3) formation of mature appressoria, (4) penetration of the cuticle and epidermal cells, (5) formation of haustoria and (6) development of elongating secondary hyphae. Each stage differed in its requirement for temperature, relative humidity and light [S, IO]. Under optimal conditions over 75% of the parasites moved through each stage with a high degree of synchrony t-6, 8, 101. Small changes in rates of s2P and s% transfer from wheat to E. graminis were detected during primary infection [9]. The amount of 32P and ssS transferred to the fungus was correlated with the development of haustoria. Differences in 9% transfer rates between compatible and incompatible parasite-host genotypes were also demonstrated [I.!?]. The increased rate of 35s transfer following 20 h after inoculation reported earlier [9, 131 was largely attributed to increased amounts of s5S being available for transfer, due to environmental influence at these times rather than to an actual increase in s5S transfer rate [15]. The time of the highest rate of transfer was found to be earlier than previously reported [9, 131 but the same differences in a5S transfer rates were found between compatible and incompatible parasite-host genotypes. t Present address of senior author: 0048-4059/78/0701~001
$02.00/O
Fort Hays Branch Experiment
Station, Hays, Kansas 67601.
@ 1978 Academic Press Inc. (London)
Limited
2
T. J. Martin and A. H. Ellingboe
If the rate of saP transfer from host to parasite were known for both compatible and incompatible parasite-host genotypes, we could correlate the transfer rates with morphological development of the parasite and also with the rates of s% transfer during the same period. Such correlations could provide a basis for a complex biochemical study on the changes in % and ssP labeled compounds during primary infection with both compatible and incompatible parasite-host genotypes. Such a study might identify a sequence of events critical to the establishment of compatible or incompatible relationships between host and parasite. The objectives of the research reported herein were : (1) to examine the uptake and translocation of 82P by the host under the environmental conditions used for synchronous parasite development, (2) to determine the rates of s2P transfer from host to parasite by considering the amount of label (theoretically available) in the epidermis for transfer to the parasite and (3) to correlate the rate of 82P transfer with the morphological stage of development of the parasite and the rates of % transfers for both compatible and incompatible parasite/host genotypes. MATERIALS AND METHODS Culture of host pathogen Inocula were obtained from cultures MS-l of E. graminis f. sp. tritici maintained on wheat ( Triticum aestiuum cv. Little Club). The cultures and environmental conditions used were as described by Masri and Ellingboe [S, 71. Plants used in experiments were 5 to 6 days old. They were uniformly inoculated by dusting with conidia from stock cultures [15]. Erwironmental conditionsfor experiments All experiments were done in Sherer-Gillett (Model CEL 512-37 and Model CEL 25-7) growth chambers. Conditions used to obtain high infection efficiency and synchronous growth of the parasite during primary infection have been reported [S, 8, 101. Light intensities used during experiments were 1-O x lo6 ergs cm-s s-1 radiation (0.6 x lo5 ergs cm -2 s-l from white VHO-fluorescent tubes and 0.4 x lo5 ergs crnm2 s-1 from 25 W incandescent bulbs). Light intensity was measured at the leaf tips with a YSI Kettering Model 65 radiometer. Designation of genotypes Briggle’s terminology [Z, 31 was used to designate the R genes conditioning reaction to E. graminis f. sp. t&i&. Genes at distinct loci have been designated Pml, Pm2, etc. Genes at the same locus but thought to be allelic are followed by the letters a, 6, c, etc. Since the host lines are all homozygous, the genotypes were written as haploid (Pmw) rather than in the diploid from (Pmu Pnrc). Detection of saP in leaf sections, epidermis and parasite The amount of *sP taken up by the leaves was determined by macerating 1 cm long leaf sections, 1 cm from the tip of each of the four leaves, in 5 ml of O-1 M NaKPO, buffer in a glass tissue grinder. A O-1 ml aliquot was dried on a piece of Whatman
‘*P transfer from wheat to Erysiphe graminis
3
no. 1 filter paper. It was then submerged in scintillation fluid [5 g of 2,5-diphenyloxazole and 0.1 g of 1,4-bis-2-(5-phenyloxazolyl) benzene in 1 1 toluene] and radioactivity was determined in a Beckman LS-133 liquid scintillation spectrometer. The data are expressed as ct/min in one leaf section. Unless otherwise stated this last method was used in all experiments. The amount of s2P translocated to the epidermis was determined by stripping approximately 50% of the epidermis from the underside of one leaf section. All green tissue was removed before placing the epidermis in scintillation vials. Scintillation fluid was added and the radioactivity determined. The data are expressed as cts/min in the abaxial epidermis of a 1 cm long leaf section. The following procedure was used to separate the ectoparasitic portion of the parasite from the leaf. A 1.9% solution of parlodion in ether : ethanol (40 : 60) was applied to a 1 cm section of a leaf [9]. The solvent evaporated in minutes and the parlodion film was easily removed with forceps. The ectoparasitic portion of the parasite was embedded within the plastic strip. Four plastic strips were dried, placed in scintillation fluid and the radioactivity determined as described above. Environmental efects of SP uptake and translocation Twenty hours after inoculation, wheat seedlings were cut at the crown with a razor blade (a film of water was present on the cutting edge) and placed into small vials (5 x 25 nm) that contained 0.1 ml of a 50 yCi/ml solution of HsS2P0, in 0.1 M NaKPO, buffer (pH 6.9). One set of plants was kept in the dark and another in the light. Temperature and relative humidity were constant. After 1, 2, 3,4, 5, 6, 8 and 10 h in s2P solution, the radioactivity in 1 cm length leaf sections was measured. The amount of 82P taken up by the leaf and translocated to the epidermis was determined under conditions necessary to induce synchronous parasite development. Inoculated wheat seedlings were cut and placed in the s2P solution. Wheat leaves were given 5 h s2P uptake periods beginning at various hours after inoculation. Different plants were used for each 5 h uptake period. Thus the amount of s2P in a leaf section at 10 h after inoculation was the result of uptake from 5 to 10 h after inoculation and the amount of s2P in a leaf section at 20 h was the result of uptake from 15 to 20 h, etc. Leaf sections and epidermal strips were taken from different plants every 2 h beginning at 6 h and ending at 30 h after inoculation. 82P transferfrom host to parasite Four parlodion strips were taken from inoculated plants which had taken up ssP for 5 h preceding the application of the parlodion. These were placed in a single vial and the amount of radioactivity determined. The results were plotted as radio activity per 5000 spores applied to the leaves. The number of spores applied was determined by making direct counts of parlodion strips made from plants inoculated at the same time but not labeled with s2P, The radioactivity in parlodion strips made from non-inoculated leaves given 82P during the same period as inoculated leaves was subtracted from the radioactivity transferred to the parasites. The rates of s2P transfer during primary infection were determined for one compatible and six incompatible parasite-host interactions. These data were then corrected for the amount of s2P theoretically available for transfer, (ct/min)/4500 ct/min in the
T. J. Martin and A. H. Ellingboe
4
epidermis, and the amount transferred per parasite unit which formed elongated secondary hyphae. Rejlication and statistics Experiments were repeated on five different days with two replications daily. The data are presented as averages of all replications. Statistical analyses were done using a two-way analysis of variance, with an F-test at the 5O/, level of significance. RESULTS
saP uptake and translocation There was a definite effect of light on s2P uptake (Fig. 1). After 5 h in the s2P solution, leaves kept in the dark were only 50% as radioactive as plants kept in the light (Fig. 1). The rate of szP uptake during this 10 h period was constant both for plants in the light and the dark.
n ‘0
2 $ ‘3
2.0 ’ I.5
”
I
I
I
I
I
I
I
2
3
4
6
6
I IO
I
Time (h) FIG. 1. The effect of light on aaP uptake in inoculated wheat leaves through being excised and placed in @P solution. (saP ct/min x 10~~ cm-l leaf section.) o, dark.
10 h after
l , Light;
The environment required to induce synchronous parasite development also influenced the uptake and translocation of s2P (Fig. 2). Both curves show a major effect of light on ssP uptake or translocation. The radioactivity in the leaf sections of plants taking up 32P from 21 to 26 h after inoculation (light period) was four times greater than in the leaf sections of those plants that took up 32P from 15 to 20 h (dark period). There was an eight-fold difference in radioactivity in the epidermis at the same uptake times. Since all of the 32P in the epidermis is theoretically available for transfer to the parasite, there were large differences in the amount of ssP
“P transfer
5
from wheat to Erysiphe graminis
available for transfer to the parasite, depending during the saP uptake period.
on the environmental
conditions
Transfer of szP to the parade The rates of szP transfer from near-isogenic wheat lines (Fig. are incompatible with culture compatible with culture MS-1
wheat leaves to E. graminis was determined, using 3). Wheat lines containing Pml, Pm2, Pm3b and Pm4 MS-1 (Px) while Chancellor, which contains pmx, is (Px). The radioactivity in parlodion strips from non-
8 5 ii
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f f n\ lo
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: ; % .
r f t
I.O-
a. : 0.5 -
I 0
I6
I
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I I4
I
IIll
I8
I
22
26
l
II
30°
after inoculation (h)
FIG. 2. 32P ct/min in wheat leaves and their epidermis under environmental conditions used for synchronous Etysi/h gruminis f. sp. tn’tici development during primary infection. 0, Leaf sections; l , epidermis.
inoculated plants which had taken up 32P during the same time was subtracted from the amount of ssP in parlodion strips from inoculated plants. This control averaged 53 to 81 ct/min. 3sP transfer from all wheat lines to fungus started at 10 h after inoculation and increased very slowly until 20 h after inoculation. There were no differences between near-isogenic lines during this period. With the compatible genotype (pmx), the rates increased sharply after 20 h, then leveled off or decreased slightly after 26 h. The rates of transfer from Pm1 were significantly less compared to pmx by 24 h. Transfer rates continued to decrease from 22 to 30 h. The transfer kinetics were the same from Pm2 and pmx. Rates of transfer from Pm3b were not significantly different from pmx until 30 h after inoculation. Transfer rates from Pm4 were similar to pmx until 28 h after inoculation, when the rate dropped rapidly. The kinetics of ssP transfer from pmx and three alleles of the Pm3 locus were determined (Fig. 4). Pm3a and Pm36 had similar rates of transfer, and were significantly lower thanpmx and Pm3c 30 h after inoculation. Pm3c did not differ frompmx.
6
T. J. Martin and A. H. Ellingboe
The rates of s2P transfer shown in Fig. 3 were adjusted for the amount of BP theoretically available in the epidermis for transfer to the parasite (Fig. 2) at each hour after inoculation for each host line (Fig. 5). The data were plotted as ssP
flme
after
inoculation
(h)
FIG. 3. Rates of srP transfer from five near-isoge.nic wheat lies to E. graminis culture MS-l (Px) during primary infection. CP et/minx 106/5000 spores applied.) 0, pmw; A, Pml;
l , Pm2; A, Pm3b; q , Pm4.
2.01
Time after
inoculation (h)
FIG. 4. Rates of srP transfer from near-kogenic
wheat lies
E. graminis culture MS-l (Px) during primary infection. applied.) 0, Pmx; A, Pmf; A, Pm3b; 0, Pm%.
of the Pm3 allelic series to
(=P ct/min x 1O-8/5OOO spores
nP transfer from wheat to Erysiphe graminis
7
(ct/min)/5000 spores/4500 ct/min available in the epidermis at the end of the 5 h uptake period. Rates of s2P transfer from the five near-isogenic lines from 6 to 20 h were not significantly different from each other, thus, these data were averaged
Time after inoculation ( h) FIG. 5. Rates of a*P transfer from five near-isogenic wheat lines to E. graminis culture MS-1 (Px) during primary infection. (9aP ct/min x 5000 spores applied/4500 “P ct/min in the epidermis.) 0, pm*; A, Pml; 0, Pm2; A, PmJ; 0, Pm4.
together and represented by a single line. With the adjustments made (Fig. 5), the same differences between host lines which were shown in Fig. 3 are seen again, but the maximum rate of transfer from pmx now appears to occur from 20 to 22 h instead of 24 to 26 h after inoculation. A steady increase in transfer rate occurred from 10 to 22 h after inoculation followed by a gradual decrease after 22 h. The rates of s2P transfer from pmx in the light and dark from 20 to 30 h after inoculation were determined (Fig. 6). The data were plotted as (ct/min)/5000 spores applied. No correction for the amount of *2P in the epidermis was made. Transfer rates in the light were similar to those reported in Fig. 3. Transfer rates in the dark at 23 and 29 h were not different from the rate of s2P transfer at 20 h. This gives support to the argument that the adjustment of transfer data to take into consideration the amount of SP activity in the epidermis is reasonable. If we assume that only functional parasite units [I,?], i.e. those with haustoria and ESH, are taking up s2P from the leaf we can further adjust the data in Fig. 4 to give the rates of 82P transfer/functional parasite unit/(ct/min) available in the epidermis (Fig. 7). Infection efficiencies on pmx, Pml, Pm36 and Pm4 were 80, 17, 30 and 4%, respectively [I.!?]. The adjustments indicated that the rates of =P transfer/successful primary infection were higher with incompatible genotypes than with compatible genotypes. At 30 h after inoculation the rates of 32P transfer/
T. J. Martin and A. Ii. Ellingboe
2.0 0 ‘px 0
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after
~.
I
22
I
26
I
30
inoculation (h)
FIG. 6. Rates of a*P transfer from Chancellor wheat (pmr) to E. gramirk culture MS-l (Px) during primary infection. Dark from 6 to 20 h and light from 20 to 30 h after inoculation (-), dark from 6 to 30 h after inoculation (---). (*sP et/minx 1O-a/5OOOspores applied.)
Time
after
Inoculation (h)
FIG. 7. Rates of ssP transfer from four near-isogenic wheat lines to E. graminis culture MS-1 (Px) during primary infection. (s*P ct/min x 10-s/5000 functional parasite units/4500 ssP ct/min in the epidermis.) 0, pmx; A, Pml; A, Pm3b; 0, Pm4.
rrP transfer from wheat to Erysiphe graminis
9
successful primary infection with the incompatible interactions with Pm1 and Pm3b were similar to the rate for the compatible interaction with pmu. Transfer rates from Pm4 were significantly different from those with pmx at 30 h, but they were decreasing rapidly. DISCUSSION
In preliminary experiments, the environmental conditions necessary for synchronous development of E. graminis f. sp. tritici were shown to influence the uptake of ssP during primary infection (Fig. 2). It appears that light is the major factor which increases transpiration, thereby increasing the uptake rate of ssP [5J. The rate of s2P uptake by plants in the dark is only about 50% of the rate of uptake by plants in the light. Earlier work [S] indicated that leaves given s2P for 4 h were saturated and additional uptake time would not greatly increase the amount of 32P taken into the leaf. It was argued that if the leaf was already saturated with s2P, the environment should not affect the amount of ssP in the leaf. However, the data presented here (Fig. 1) conclusively show that light does affect s2P uptake and saturation does not occur for at least 10 h. Earlier experiments were done with 100 @ i s2P/ml solutions; 50 &i/ml was used in this study. This difference should have been negligible because the s2P was in a 0.1 M phosphate buffer in both studies. The differences between earlier studies and the data presented herein may be due to improved procedures which minimize the introduction of air into the vascular system when leaves were cut at the base and placed in solutions containing 32P. If YS and s2P were both transferred from host to parasite by simple diffusion, one would expect similar rates of transfer for each isotope. However, there are some important differences between the data reported in this paper for transfer of s2P and the previously reported data on ?S transfer for a compatible parasite-host interaction [15]. The earliest time at which 32P was detected in the parasite was 10 h after inoculation, which corresponds to the time at which haustoria can first be seen in the host cells [ 81. 3sS was first detected in the parasite at 16 h, which is the time at which haustoria begin to develop appendages [4]. This may indicate that ssP simply diffuses into the parasite as soon as the parasite penetrates the host cell wall, while YS may require a more complex transport system. On the other hand, it may be that the transport systems for 32P function earlier than those for 85s. s2P and 35S transfer rates were very similar from 18 h onward. However, the amount of 32P transferred was much higher. This could be a function of the ability of ssP to move through the leaf faster than 35s. When corrections were applied for the amount of s2P available for transfer, the highest rate of transfer occurred from 20 to 22 h after inoculation. Similar results were obtained for 55s transfer [15]. The rates of s2P and s% transfer decreased after 22 h. The validity of correcting transfer rates for the amount of s2P available is supported by the fact that when the lights were left off from 20 to 30 h, the amount of 32P available for transfer remained constant. The dark period from 20 to 30 h does slow down development of ESH but only for 2 h [14]. Thus, slower rates of ESH development would not explain the decreased transfer in the dark. The amount of s2P transferred from host to parasite was found to correspond to morphological development of the fungus with the various genotypes. The
10
T. J. Martin and A. H. Ellingboe
reduction of the percentage of ESH for the incompatible interactions [12], PlIPml, P2/pm2, P3bjPm36, and P4/Pm4, was similar to the reduction in s2P transfer, however, the decreases in s2P transfer were later than those observed for % transfer. Decreases in 32P transfer were not seen until after morphological differences were noted. Y+l transfer was affected concurrently with or prior to effects of the different incompatible genotypes on morphological development of the fungus [15]. Results from the comparison of s2P transfer from host lines with the allelic series of genes at the Pm3 locus failed to differentiate between Pm3a and Pm3b. If these genes are truly allelic, then one would expect them to affect parasite development in the same way. Pm3c had similar kinetics of transfer to jmx. This was expected since Pm3c does not have an apparent effect in the primary leaf [I, 2, II]. If only functional parasite units transfer 82P, then the amount of 32P transferred by each functional unit in incompatible interactions is greater than that transferred in compatible interactions. With the exception of Pm4, the rates of ssP transfer in incompatible interactions eventually decreased to the same level as in the compatible interactions. Rates of s2P transfer from plants with Pm4 were higher at 30 h, but they were decreasing and may have eventually reached the same as in the compatible interactions. An alternate explanation is that the non-functional parasite units could be taking up a2P until 30 h after inoculation, at which time the process is halted. It is also possible, especially with P4 and Pm4, that, after the collapse of the parasite, which occurs approximately 22 h after inoculation, there was diffusive flow of s2P into the collapsed parasite. By 30 h after inoculation, this “dead” parasite may have been in a condition such that S2P did not diffuse into it or the necrotic host cell which the parasite had attempted to penetrate. The data reported here along with previously published data on ssS transfer and morphological development during primary infection [S, 10, 1.51 lay a solid base for more detailed biochemical studies. Careful study of the changes in YS and ssaPlabelled compounds during primary infection may contribute to the elucidation of a sequence of events critical to the establishment of compatible or incompatible relationships between host and parasite. This research was supported in part by Grant A106420 from the National Institute of Health and Grant GB41214 from the National Science Foundation. Michigan Agricultural Experiment Station, Journal Article No. 8324. REFERENCES 1. BRIGGLE, L. W. & SCHAREN, A. L. (1961). Resistance in Tritkwn vulgare to infection by Erysiph gruminis f. sp. titici as influenced by the stage of development of the host plant. Plant Direas RcportH 45,846-850. 2. BRIGGLJI, L. W. (1966). Three loci in wheat involving resistance to Erys+Vu graminis f. sp. tritici. Crop Science6, 461-465. 3. BIUCGLE, L. W. (1969). Near-isogenic lines of wheat with genes for resistance to Erpsiphc graminis f. sp. tritici. crop s&ncc 9,70-75. 4. IIAYWOOD, M. J., SR (1975). Genetic control of the development of haustoria of E~si@u graminis f. sp. tdici on wheat. Ph.D. Thesis, Michigan State University. 5. MARTIN, T. J., STUCKEY, R. E., Sm, G. R. & ELLINGBOE, A. H. (1975). Reduction of transpiration from wheat caused by germinating conidii of E&he graminis f. sp. tritici. PhysioIogical Plant Pathology 7, 71-77.
=P transfer from wheat to Erysiphe graminis
11
6. MASRI, S. S. & ELLINCBOE, A. H. (1966). Germination of conidia and formation of appressoria and secondary hyphae in Erysi#w graminis f. ap. triti. Phyto@thology 56, 304-308. 7. MASRI, S. S. & ELLINGBOE, A. H. (1966). Primary infection of wheat and barley by Ep$#u graminis. Phytojdzology 56,38!&395. 8. MOUNT, M. S. (1968). Environmental effects and transfer events during primary infection of wheat by Esvsiphc graminis. Ph.D. Thesis, Michigan State University. 9. MOUNT, M. S. & ELLINGBOE, A. H. (1969). sap and W transfer from susceptible wheat to Etysiphc graminis f. ap. tritici during primary infection. Phyto#athology59, 235. 10. NAIR, K. R. S. & ELLINGBOE, A. H. (1965). Germination of conidia of Etysifihe graminis f. sp. tritici. Phytopathology55, 365-368. 11. PUGSLEY,A. T. & CARTER, M. V. (1953). The resistance of twelve varieties of ~&cum uulgare to [email protected] gramiais tritici. Australian Journalof Biological Sciences6,335-346. 12. SLESINSKI,R. S. & ELLINGBOE, A. H. (1969). The genetic control of primary infection of wheat by Ery$he graminis f. ap. tritici. Phyropatilogv 59,1833-1837. 13. SLESINSIU,R. S. & ELLINCBOE, A. H. (1971). Transfer of ssS from wheat to the powdery mildew fungus with compatible and incompatible parasite/host genotypes. Canadian journal of Botany 49,303~310. 14. STUCKEY, R. E. (1973). Elongation of secondary hyphae and translocation of 8% and *H from host to parasite during primary infection of wheat by E&.he graminis f. ap. t&id. Ph.D. Thesis, Michigan State University. 15. STUCKEY, R. E. & ELLINQBOE, A. H. (1975). Effect of environmental conditions on ssS uptake by Triticum aestimnn and transfer to Erysiphe graminis f. sp. tritici during primary infection. Physiological Plant Pathology 5, 19-26.
Genetic control of 32Ptransfer from wheat to Erysiphe gram inis f. sp. frifici during primary infection T. J. MARTIN? and A. H. ELLINGBOE Departmentof Botany and Plant Pathology, Michigan State University East Lansing, Michigan 48824, U.S.A. (AcceptedforpublicationJanuary 1978)
Environmental conditions necessary for synchronous development of Etysiphe graminis D.C. f. sp. tritici Em. Marchal during primary infection of wheat were shown to affect the amount of szP taken up and translocated to the epidermis of excised leaves placed in a saP solution. Light increased transpiration, the amount of 32P taken up by the leaves and the amount of s2P transferred to the parasite. Rates of saP transfer from host to parasite were determined for compatible and incompatible parasite-host interactions during primary infection. Transfer rate reflected the relative compatibility of the host-parasite interaction studied. Interactions with little fungal development had lower total rates of transfer than did compatible interactions. However, when transfer was expressed as saP transferred per successful primary infection, the successful parasite units involved in incompatible interactions obtained saP at a greater rate than did parasite units in compatible interactions.
INTRODUCTION
The primary infection process of Erysiphe graminis on wheat (T&cum aestivum) has been divided into distinct morphological stages : (1) germination, (2) production of club-shaped appressorial initials, (3) formation of mature appressoria, (4) penetration of the cuticle and epidermal cells, (5) formation of haustoria and (6) development of elongating secondary hyphae. Each stage differed in its requirement for temperature, relative humidity and light [S, IO]. Under optimal conditions over 75% of the parasites moved through each stage with a high degree of synchrony t-6, 8, 101. Small changes in rates of s2P and s% transfer from wheat to E. graminis were detected during primary infection [9]. The amount of 32P and ssS transferred to the fungus was correlated with the development of haustoria. Differences in 9% transfer rates between compatible and incompatible parasite-host genotypes were also demonstrated [I.!?]. The increased rate of 35s transfer following 20 h after inoculation reported earlier [9, 131 was largely attributed to increased amounts of s5S being available for transfer, due to environmental influence at these times rather than to an actual increase in s5S transfer rate [15]. The time of the highest rate of transfer was found to be earlier than previously reported [9, 131 but the same differences in a5S transfer rates were found between compatible and incompatible parasite-host genotypes. t Present address of senior author: 0048-4059/78/0701~001
$02.00/O
Fort Hays Branch Experiment
Station, Hays, Kansas 67601.
@ 1978 Academic Press Inc. (London)
Limited
2
T. J. Martin and A. H. Ellingboe
If the rate of saP transfer from host to parasite were known for both compatible and incompatible parasite-host genotypes, we could correlate the transfer rates with morphological development of the parasite and also with the rates of s% transfer during the same period. Such correlations could provide a basis for a complex biochemical study on the changes in % and ssP labeled compounds during primary infection with both compatible and incompatible parasite-host genotypes. Such a study might identify a sequence of events critical to the establishment of compatible or incompatible relationships between host and parasite. The objectives of the research reported herein were : (1) to examine the uptake and translocation of 82P by the host under the environmental conditions used for synchronous parasite development, (2) to determine the rates of s2P transfer from host to parasite by considering the amount of label (theoretically available) in the epidermis for transfer to the parasite and (3) to correlate the rate of 82P transfer with the morphological stage of development of the parasite and the rates of % transfers for both compatible and incompatible parasite/host genotypes. MATERIALS AND METHODS Culture of host pathogen Inocula were obtained from cultures MS-l of E. graminis f. sp. tritici maintained on wheat ( Triticum aestiuum cv. Little Club). The cultures and environmental conditions used were as described by Masri and Ellingboe [S, 71. Plants used in experiments were 5 to 6 days old. They were uniformly inoculated by dusting with conidia from stock cultures [15]. Erwironmental conditionsfor experiments All experiments were done in Sherer-Gillett (Model CEL 512-37 and Model CEL 25-7) growth chambers. Conditions used to obtain high infection efficiency and synchronous growth of the parasite during primary infection have been reported [S, 8, 101. Light intensities used during experiments were 1-O x lo6 ergs cm-s s-1 radiation (0.6 x lo5 ergs cm -2 s-l from white VHO-fluorescent tubes and 0.4 x lo5 ergs crnm2 s-1 from 25 W incandescent bulbs). Light intensity was measured at the leaf tips with a YSI Kettering Model 65 radiometer. Designation of genotypes Briggle’s terminology [Z, 31 was used to designate the R genes conditioning reaction to E. graminis f. sp. t&i&. Genes at distinct loci have been designated Pml, Pm2, etc. Genes at the same locus but thought to be allelic are followed by the letters a, 6, c, etc. Since the host lines are all homozygous, the genotypes were written as haploid (Pmw) rather than in the diploid from (Pmu Pnrc). Detection of saP in leaf sections, epidermis and parasite The amount of *sP taken up by the leaves was determined by macerating 1 cm long leaf sections, 1 cm from the tip of each of the four leaves, in 5 ml of O-1 M NaKPO, buffer in a glass tissue grinder. A O-1 ml aliquot was dried on a piece of Whatman
‘*P transfer from wheat to Erysiphe graminis
3
no. 1 filter paper. It was then submerged in scintillation fluid [5 g of 2,5-diphenyloxazole and 0.1 g of 1,4-bis-2-(5-phenyloxazolyl) benzene in 1 1 toluene] and radioactivity was determined in a Beckman LS-133 liquid scintillation spectrometer. The data are expressed as ct/min in one leaf section. Unless otherwise stated this last method was used in all experiments. The amount of s2P translocated to the epidermis was determined by stripping approximately 50% of the epidermis from the underside of one leaf section. All green tissue was removed before placing the epidermis in scintillation vials. Scintillation fluid was added and the radioactivity determined. The data are expressed as cts/min in the abaxial epidermis of a 1 cm long leaf section. The following procedure was used to separate the ectoparasitic portion of the parasite from the leaf. A 1.9% solution of parlodion in ether : ethanol (40 : 60) was applied to a 1 cm section of a leaf [9]. The solvent evaporated in minutes and the parlodion film was easily removed with forceps. The ectoparasitic portion of the parasite was embedded within the plastic strip. Four plastic strips were dried, placed in scintillation fluid and the radioactivity determined as described above. Environmental efects of SP uptake and translocation Twenty hours after inoculation, wheat seedlings were cut at the crown with a razor blade (a film of water was present on the cutting edge) and placed into small vials (5 x 25 nm) that contained 0.1 ml of a 50 yCi/ml solution of HsS2P0, in 0.1 M NaKPO, buffer (pH 6.9). One set of plants was kept in the dark and another in the light. Temperature and relative humidity were constant. After 1, 2, 3,4, 5, 6, 8 and 10 h in s2P solution, the radioactivity in 1 cm length leaf sections was measured. The amount of 82P taken up by the leaf and translocated to the epidermis was determined under conditions necessary to induce synchronous parasite development. Inoculated wheat seedlings were cut and placed in the s2P solution. Wheat leaves were given 5 h s2P uptake periods beginning at various hours after inoculation. Different plants were used for each 5 h uptake period. Thus the amount of s2P in a leaf section at 10 h after inoculation was the result of uptake from 5 to 10 h after inoculation and the amount of s2P in a leaf section at 20 h was the result of uptake from 15 to 20 h, etc. Leaf sections and epidermal strips were taken from different plants every 2 h beginning at 6 h and ending at 30 h after inoculation. 82P transferfrom host to parasite Four parlodion strips were taken from inoculated plants which had taken up ssP for 5 h preceding the application of the parlodion. These were placed in a single vial and the amount of radioactivity determined. The results were plotted as radio activity per 5000 spores applied to the leaves. The number of spores applied was determined by making direct counts of parlodion strips made from plants inoculated at the same time but not labeled with s2P, The radioactivity in parlodion strips made from non-inoculated leaves given 82P during the same period as inoculated leaves was subtracted from the radioactivity transferred to the parasites. The rates of s2P transfer during primary infection were determined for one compatible and six incompatible parasite-host interactions. These data were then corrected for the amount of s2P theoretically available for transfer, (ct/min)/4500 ct/min in the
T. J. Martin and A. H. Ellingboe
4
epidermis, and the amount transferred per parasite unit which formed elongated secondary hyphae. Rejlication and statistics Experiments were repeated on five different days with two replications daily. The data are presented as averages of all replications. Statistical analyses were done using a two-way analysis of variance, with an F-test at the 5O/, level of significance. RESULTS
saP uptake and translocation There was a definite effect of light on s2P uptake (Fig. 1). After 5 h in the s2P solution, leaves kept in the dark were only 50% as radioactive as plants kept in the light (Fig. 1). The rate of szP uptake during this 10 h period was constant both for plants in the light and the dark.
n ‘0
2 $ ‘3
2.0 ’ I.5
”
I
I
I
I
I
I
I
2
3
4
6
6
I IO
I
Time (h) FIG. 1. The effect of light on aaP uptake in inoculated wheat leaves through being excised and placed in @P solution. (saP ct/min x 10~~ cm-l leaf section.) o, dark.
10 h after
l , Light;
The environment required to induce synchronous parasite development also influenced the uptake and translocation of s2P (Fig. 2). Both curves show a major effect of light on ssP uptake or translocation. The radioactivity in the leaf sections of plants taking up 32P from 21 to 26 h after inoculation (light period) was four times greater than in the leaf sections of those plants that took up 32P from 15 to 20 h (dark period). There was an eight-fold difference in radioactivity in the epidermis at the same uptake times. Since all of the 32P in the epidermis is theoretically available for transfer to the parasite, there were large differences in the amount of ssP
“P transfer
5
from wheat to Erysiphe graminis
available for transfer to the parasite, depending during the saP uptake period.
on the environmental
conditions
Transfer of szP to the parade The rates of szP transfer from near-isogenic wheat lines (Fig. are incompatible with culture compatible with culture MS-1
wheat leaves to E. graminis was determined, using 3). Wheat lines containing Pml, Pm2, Pm3b and Pm4 MS-1 (Px) while Chancellor, which contains pmx, is (Px). The radioactivity in parlodion strips from non-
8 5 ii
2-o-
f f n\ lo
1.5-
: ; % .
r f t
I.O-
a. : 0.5 -
I 0
I6
I
IO Time
I I4
I
IIll
I8
I
22
26
l
II
30°
after inoculation (h)
FIG. 2. 32P ct/min in wheat leaves and their epidermis under environmental conditions used for synchronous Etysi/h gruminis f. sp. tn’tici development during primary infection. 0, Leaf sections; l , epidermis.
inoculated plants which had taken up 32P during the same time was subtracted from the amount of ssP in parlodion strips from inoculated plants. This control averaged 53 to 81 ct/min. 3sP transfer from all wheat lines to fungus started at 10 h after inoculation and increased very slowly until 20 h after inoculation. There were no differences between near-isogenic lines during this period. With the compatible genotype (pmx), the rates increased sharply after 20 h, then leveled off or decreased slightly after 26 h. The rates of transfer from Pm1 were significantly less compared to pmx by 24 h. Transfer rates continued to decrease from 22 to 30 h. The transfer kinetics were the same from Pm2 and pmx. Rates of transfer from Pm3b were not significantly different from pmx until 30 h after inoculation. Transfer rates from Pm4 were similar to pmx until 28 h after inoculation, when the rate dropped rapidly. The kinetics of ssP transfer from pmx and three alleles of the Pm3 locus were determined (Fig. 4). Pm3a and Pm36 had similar rates of transfer, and were significantly lower thanpmx and Pm3c 30 h after inoculation. Pm3c did not differ frompmx.
6
T. J. Martin and A. H. Ellingboe
The rates of s2P transfer shown in Fig. 3 were adjusted for the amount of BP theoretically available in the epidermis for transfer to the parasite (Fig. 2) at each hour after inoculation for each host line (Fig. 5). The data were plotted as ssP
flme
after
inoculation
(h)
FIG. 3. Rates of srP transfer from five near-isoge.nic wheat lies to E. graminis culture MS-l (Px) during primary infection. CP et/minx 106/5000 spores applied.) 0, pmw; A, Pml;
l , Pm2; A, Pm3b; q , Pm4.
2.01
Time after
inoculation (h)
FIG. 4. Rates of srP transfer from near-kogenic
wheat lies
E. graminis culture MS-l (Px) during primary infection. applied.) 0, Pmx; A, Pmf; A, Pm3b; 0, Pm%.
of the Pm3 allelic series to
(=P ct/min x 1O-8/5OOO spores
nP transfer from wheat to Erysiphe graminis
7
(ct/min)/5000 spores/4500 ct/min available in the epidermis at the end of the 5 h uptake period. Rates of s2P transfer from the five near-isogenic lines from 6 to 20 h were not significantly different from each other, thus, these data were averaged
Time after inoculation ( h) FIG. 5. Rates of a*P transfer from five near-isogenic wheat lines to E. graminis culture MS-1 (Px) during primary infection. (9aP ct/min x 5000 spores applied/4500 “P ct/min in the epidermis.) 0, pm*; A, Pml; 0, Pm2; A, PmJ; 0, Pm4.
together and represented by a single line. With the adjustments made (Fig. 5), the same differences between host lines which were shown in Fig. 3 are seen again, but the maximum rate of transfer from pmx now appears to occur from 20 to 22 h instead of 24 to 26 h after inoculation. A steady increase in transfer rate occurred from 10 to 22 h after inoculation followed by a gradual decrease after 22 h. The rates of s2P transfer from pmx in the light and dark from 20 to 30 h after inoculation were determined (Fig. 6). The data were plotted as (ct/min)/5000 spores applied. No correction for the amount of *2P in the epidermis was made. Transfer rates in the light were similar to those reported in Fig. 3. Transfer rates in the dark at 23 and 29 h were not different from the rate of s2P transfer at 20 h. This gives support to the argument that the adjustment of transfer data to take into consideration the amount of SP activity in the epidermis is reasonable. If we assume that only functional parasite units [I,?], i.e. those with haustoria and ESH, are taking up s2P from the leaf we can further adjust the data in Fig. 4 to give the rates of 82P transfer/functional parasite unit/(ct/min) available in the epidermis (Fig. 7). Infection efficiencies on pmx, Pml, Pm36 and Pm4 were 80, 17, 30 and 4%, respectively [I.!?]. The adjustments indicated that the rates of =P transfer/successful primary infection were higher with incompatible genotypes than with compatible genotypes. At 30 h after inoculation the rates of 32P transfer/
T. J. Martin and A. Ii. Ellingboe
2.0 0 ‘px 0
1.5-
:E e ‘,
I.O-
0 /’
0 -0-o
z 0
0.5 I’ ~~.-~‘----~---‘-~_l_
0
I 6
2 l --lj/ 10
I
’
18 Titi
after
~.
I
22
I
26
I
30
inoculation (h)
FIG. 6. Rates of a*P transfer from Chancellor wheat (pmr) to E. gramirk culture MS-l (Px) during primary infection. Dark from 6 to 20 h and light from 20 to 30 h after inoculation (-), dark from 6 to 30 h after inoculation (---). (*sP et/minx 1O-a/5OOOspores applied.)
Time
after
Inoculation (h)
FIG. 7. Rates of ssP transfer from four near-isogenic wheat lines to E. graminis culture MS-1 (Px) during primary infection. (s*P ct/min x 10-s/5000 functional parasite units/4500 ssP ct/min in the epidermis.) 0, pmx; A, Pml; A, Pm3b; 0, Pm4.
rrP transfer from wheat to Erysiphe graminis
9
successful primary infection with the incompatible interactions with Pm1 and Pm3b were similar to the rate for the compatible interaction with pmu. Transfer rates from Pm4 were significantly different from those with pmx at 30 h, but they were decreasing rapidly. DISCUSSION
In preliminary experiments, the environmental conditions necessary for synchronous development of E. graminis f. sp. tritici were shown to influence the uptake of ssP during primary infection (Fig. 2). It appears that light is the major factor which increases transpiration, thereby increasing the uptake rate of ssP [5J. The rate of s2P uptake by plants in the dark is only about 50% of the rate of uptake by plants in the light. Earlier work [S] indicated that leaves given s2P for 4 h were saturated and additional uptake time would not greatly increase the amount of 32P taken into the leaf. It was argued that if the leaf was already saturated with s2P, the environment should not affect the amount of ssP in the leaf. However, the data presented here (Fig. 1) conclusively show that light does affect s2P uptake and saturation does not occur for at least 10 h. Earlier experiments were done with 100 @ i s2P/ml solutions; 50 &i/ml was used in this study. This difference should have been negligible because the s2P was in a 0.1 M phosphate buffer in both studies. The differences between earlier studies and the data presented herein may be due to improved procedures which minimize the introduction of air into the vascular system when leaves were cut at the base and placed in solutions containing 32P. If YS and s2P were both transferred from host to parasite by simple diffusion, one would expect similar rates of transfer for each isotope. However, there are some important differences between the data reported in this paper for transfer of s2P and the previously reported data on ?S transfer for a compatible parasite-host interaction [15]. The earliest time at which 32P was detected in the parasite was 10 h after inoculation, which corresponds to the time at which haustoria can first be seen in the host cells [ 81. 3sS was first detected in the parasite at 16 h, which is the time at which haustoria begin to develop appendages [4]. This may indicate that ssP simply diffuses into the parasite as soon as the parasite penetrates the host cell wall, while YS may require a more complex transport system. On the other hand, it may be that the transport systems for 32P function earlier than those for 85s. s2P and 35S transfer rates were very similar from 18 h onward. However, the amount of 32P transferred was much higher. This could be a function of the ability of ssP to move through the leaf faster than 35s. When corrections were applied for the amount of s2P available for transfer, the highest rate of transfer occurred from 20 to 22 h after inoculation. Similar results were obtained for 55s transfer [15]. The rates of s2P and s% transfer decreased after 22 h. The validity of correcting transfer rates for the amount of s2P available is supported by the fact that when the lights were left off from 20 to 30 h, the amount of 32P available for transfer remained constant. The dark period from 20 to 30 h does slow down development of ESH but only for 2 h [14]. Thus, slower rates of ESH development would not explain the decreased transfer in the dark. The amount of s2P transferred from host to parasite was found to correspond to morphological development of the fungus with the various genotypes. The
10
T. J. Martin and A. H. Ellingboe
reduction of the percentage of ESH for the incompatible interactions [12], PlIPml, P2/pm2, P3bjPm36, and P4/Pm4, was similar to the reduction in s2P transfer, however, the decreases in s2P transfer were later than those observed for % transfer. Decreases in 32P transfer were not seen until after morphological differences were noted. Y+l transfer was affected concurrently with or prior to effects of the different incompatible genotypes on morphological development of the fungus [15]. Results from the comparison of s2P transfer from host lines with the allelic series of genes at the Pm3 locus failed to differentiate between Pm3a and Pm3b. If these genes are truly allelic, then one would expect them to affect parasite development in the same way. Pm3c had similar kinetics of transfer to jmx. This was expected since Pm3c does not have an apparent effect in the primary leaf [I, 2, II]. If only functional parasite units transfer 82P, then the amount of 32P transferred by each functional unit in incompatible interactions is greater than that transferred in compatible interactions. With the exception of Pm4, the rates of ssP transfer in incompatible interactions eventually decreased to the same level as in the compatible interactions. Rates of s2P transfer from plants with Pm4 were higher at 30 h, but they were decreasing and may have eventually reached the same as in the compatible interactions. An alternate explanation is that the non-functional parasite units could be taking up a2P until 30 h after inoculation, at which time the process is halted. It is also possible, especially with P4 and Pm4, that, after the collapse of the parasite, which occurs approximately 22 h after inoculation, there was diffusive flow of s2P into the collapsed parasite. By 30 h after inoculation, this “dead” parasite may have been in a condition such that S2P did not diffuse into it or the necrotic host cell which the parasite had attempted to penetrate. The data reported here along with previously published data on ssS transfer and morphological development during primary infection [S, 10, 1.51 lay a solid base for more detailed biochemical studies. Careful study of the changes in YS and ssaPlabelled compounds during primary infection may contribute to the elucidation of a sequence of events critical to the establishment of compatible or incompatible relationships between host and parasite. This research was supported in part by Grant A106420 from the National Institute of Health and Grant GB41214 from the National Science Foundation. Michigan Agricultural Experiment Station, Journal Article No. 8324. REFERENCES 1. BRIGGLE, L. W. & SCHAREN, A. L. (1961). Resistance in Tritkwn vulgare to infection by Erysiph gruminis f. sp. titici as influenced by the stage of development of the host plant. Plant Direas RcportH 45,846-850. 2. BRIGGLJI, L. W. (1966). Three loci in wheat involving resistance to Erys+Vu graminis f. sp. tritici. Crop Science6, 461-465. 3. BIUCGLE, L. W. (1969). Near-isogenic lines of wheat with genes for resistance to Erpsiphc graminis f. sp. tritici. crop s&ncc 9,70-75. 4. IIAYWOOD, M. J., SR (1975). Genetic control of the development of haustoria of E~si@u graminis f. sp. tdici on wheat. Ph.D. Thesis, Michigan State University. 5. MARTIN, T. J., STUCKEY, R. E., Sm, G. R. & ELLINGBOE, A. H. (1975). Reduction of transpiration from wheat caused by germinating conidii of E&he graminis f. sp. tritici. PhysioIogical Plant Pathology 7, 71-77.
=P transfer from wheat to Erysiphe graminis
11
6. MASRI, S. S. & ELLINCBOE, A. H. (1966). Germination of conidia and formation of appressoria and secondary hyphae in Erysi#w graminis f. ap. triti. Phyto@thology 56, 304-308. 7. MASRI, S. S. & ELLINGBOE, A. H. (1966). Primary infection of wheat and barley by Ep$#u graminis. Phytojdzology 56,38!&395. 8. MOUNT, M. S. (1968). Environmental effects and transfer events during primary infection of wheat by Esvsiphc graminis. Ph.D. Thesis, Michigan State University. 9. MOUNT, M. S. & ELLINGBOE, A. H. (1969). sap and W transfer from susceptible wheat to Etysiphc graminis f. ap. tritici during primary infection. Phyto#athology59, 235. 10. NAIR, K. R. S. & ELLINGBOE, A. H. (1965). Germination of conidia of Etysifihe graminis f. sp. tritici. Phytopathology55, 365-368. 11. PUGSLEY,A. T. & CARTER, M. V. (1953). The resistance of twelve varieties of ~&cum uulgare to [email protected] gramiais tritici. Australian Journalof Biological Sciences6,335-346. 12. SLESINSKI,R. S. & ELLINGBOE, A. H. (1969). The genetic control of primary infection of wheat by Ery$he graminis f. ap. tritici. Phyropatilogv 59,1833-1837. 13. SLESINSIU,R. S. & ELLINCBOE, A. H. (1971). Transfer of ssS from wheat to the powdery mildew fungus with compatible and incompatible parasite/host genotypes. Canadian journal of Botany 49,303~310. 14. STUCKEY, R. E. (1973). Elongation of secondary hyphae and translocation of 8% and *H from host to parasite during primary infection of wheat by E&.he graminis f. ap. t&id. Ph.D. Thesis, Michigan State University. 15. STUCKEY, R. E. & ELLINQBOE, A. H. (1975). Effect of environmental conditions on ssS uptake by Triticum aestimnn and transfer to Erysiphe graminis f. sp. tritici during primary infection. Physiological Plant Pathology 5, 19-26.