INTRODUCTION

Top Abstract Introduction Materials and Methods Results Discussion References

Beet western yellows virus (BWYV) is a member of the Polerovirus genus of the Luteoviridae family (6). Luteoviruses are transmitted by aphids in a circulative nonpropagative manner and multiply in the phloem tissue of their hosts (22). The manner in which luteoviruses are acquired and transmitted by their specific vectors is complex. After uptake during feeding, the virus must first traverse the epithelial cell barrier separating the alimentary canal from the hemocoel, a process which involves receptor-mediated endocytosis and exocytosis (10, 11, 13). Once in the hemolymph, the virions diffuse until they encounter an accessory salivary gland (ASG). At this point, the virus must first penetrate the ASG basal lamina, which can act as a selective barrier to certain virus species (15, 25), and then undergo another round of receptor-mediated endocytosis and exocytosis to move into the salivary duct (13). Interactions between the virus capsid and aphid components at all of these sites may be important in conferring vector specificity (12, 14). Finally, in the hemocoel, binding of the virus to symbionin, a chaperonin secreted into the hemolymph by endosymbiotic bacteria of the aphid, is known to stabilize luteoviruses in a vector-nonspecific manner (9, 32, 33).

The genome of the poleroviruses consists of a monopartite plus-sense RNA of about 5.6 kb contained in isometric particles of 25-nm diameter (22, 23). Six major open reading frames (ORFs) are arranged on the genomic RNA into 5'-proximal (ORF0, ORF1, and ORF2) and 3'-proximal (ORF3, ORF4, and ORF5) gene clusters (see Fig. 1 for a genetic map). The 3'-located genes are expressed from a subgenomic RNA (7, 29, 35). ORF3 encodes the major capsid protein of about 22 kDa. ORF4 is embedded in the capsid protein gene in another reading frame and encodes a 17- to 19-kDa protein. In potato leafroll virus, this protein has many of the properties expected for a viral movement protein (27, 29). BWYV P19, on the other hand, is not required for whole-plant infection of several hosts (37), suggesting that another BWYV protein or proteins may substitute for or act in parallel to P19 in supplying movement functions.

View larger version (15K): [in this window] [in a new window]   FIG. 1.   Genetic map of BWYV (A) and positions of the point mutations in the RTD (B). Hatching indicates the proline tract, and stippling indicates the conserved portion of the RTD. The identity of each point mutant is indicated in bold type (5.121 = BW5.121, etc.). The amino acid(s) substituted in each mutant and the corresponding wild-type amino acid(s) are given below. Amino acid numbering begins with the first residue of the RTD. The positions of some of the restrictions sites mentioned in the text are shown at the top as B (BamHI), H (HindIII), M (MscI), N (NcoI), and S (SpeI).

The 3'-proximal ORF 5 or readthrough domain (RTD) of BWYV, as in other poleroviruses, is translated as a 74-kDa fusion protein (known as P74 or RT protein) by episodic readthrough of the major coat protein termination codon (34). The RT protein is a minor component of the luteoviral capsid (1, 2, 20, 36). The RTD consists of several subdomains (22). A cytidine-rich sequence encoding a tract of 7 to 13 alternating proline residues in the RTD is located just downstream of the coat protein cistron-suppressible termination codon. This is followed by a region of about 200 amino acid residues with considerable sequence similarity throughout the Luteoviridae. The C-terminal half of the RTD is divergent. During virus purification, the RT protein of BWYV and other luteoviruses is cleaved to produce a C-terminally truncated form of about 53 kDa (1, 2, 36).

Mutagenesis of cloned full-length luteovirus cDNA has revealed that much of the C-terminal half of the RTD is dispensable for whole-plant infection and aphid transmission (2, 4). The conserved portion of the RTD, on the other hand, contains domains which are important for (i) aphid transmission of the virus (2, 4, 5), (ii) efficient accumulation of the virus in whole plants (4, 5, 24), and (iii) efficient suppression of major coat protein translation-termination (3, 4).

In this study we have produced a set of point mutants, mostly alanine substitutions, in the conserved part of the BWYV RTD in an attempt to better characterize determinants involved in aphid transmission. The mutants were assayed by infection of protoplasts with full-length transcripts and by agroinoculation of plants, followed by aphid transmission tests using either infected protoplast extracts, agroinfected plants, or purified virus as a virus source. The stability of the various mutations was assessed by sequence analysis of the mutant progeny in the agroinfected and aphid-infected plants. The results have allowed us to identify amino acids important for virus accumulation in planta and for transmission of BWYV by Myzus persicae Sulz. Our observations also reveal that successful aphid transmission of the RTD mutants is often accompanied by compensatory mutations elsewhere in the RTD, suggesting that the RTD possesses considerable structural redundancy.

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results Discussion References

Mutants of BWYV. All point mutants were created by PCR mutagenesis (16) using pBW₀ as the template and oligonucleotides BW35 (nucleotides [nt] 4003 to 4022) and BW21 (complementary to nt 4829 to 4846) as external primers. The mutagenic primers were designed to substitute alanine residues at position 24 (mutant BW5.121; numbering refers to amino acids in the RTD), positions 59 and 60 (mutant BW5.123), positions 113 and 114 (BW5.125), and positions 225 and 226 (BW5.129). In mutant BW5.127, the amino acids K200 and Y201 were replaced by AD (Fig. 1). Full-length viral cDNAs containing the different mutations were reconstructed by cutting the PCR fragments bearing the mutations with BamHI and NcoI and substituting this fragment for the wild-type BamHI-NcoI (nt 4006 to 4822) fragment in pBW₀. The 1.1-kb BamHI fragment (nt 2904 to 4006) was then introduced to create the final full-length mutant transcription vectors.

Infection of protoplasts and plants. Protoplasts of Chenopodium quinoa were inoculated with viral RNA transcripts as described elsewhere (4). Mutant constructs for agroinfection (pBinBW5.121, pBinBW5.123, etc.) were made by replacing the SpeI-SalI fragment (extends from nt 1350 to 32 nt downstream of the insert 3' terminus) of pBinBW₀ by the SpeI-SalI fragment from the corresponding mutant transcription vector. The resulting plasmids were introduced into Agrobacterium tumefaciens LBA4404 for agroinfection (2, 18). Infected plants were identified by double-antibody sandwich enzyme-linked immunosorbent assay (DAS-ELISA) with a rabbit polyclonal antiserum raised against virus (4). Virus was purified as described by van den Heuvel et al. (31).

Aphid transmission assays. Transmission experiments used either detached leaves, crude extracts of infected protoplasts, or purified virus as an inoculum (4) and second- to fourth-instar M. persicae larvae as the vector. Membrane feeding of the larvae on crude protoplast extracts and on purified virus suspensions was performed as described elsewhere (4). For microinjection (4), the larvae were positioned with a micromanipulator and injected with 10 or 20 nl of purified virus (25 µg/ml), using 12- to 15-µm (outer diameter) glass capillaries attached to an Inject+ Matic microinjector (Gabay Instruments, Geneva, Switzerland).

Analysis of viral RNA and capsid proteins. BWYV RNA in total RNA extracts of infected plants and protoplasts was detected by Northern blotting using a ³²P-labeled RNA probe complementary to the 3'-terminal 196 nt of the viral RNA (26). Viral structural proteins in total protein extracts of BWYV-infected protoplasts and plants and in purified virus were detected by Western blotting using antisera specific for BWYV coat protein and RT protein (26) following sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) as described previously (2) except that urea was omitted from the gel loading buffer and immunodetection was performed with an enhanced chemiluminescence Western blotting kit (Amersham or Pierce).

Mutant progeny analysis. The stability of the mutations in progeny viral RNA following agroinfection or aphid transmission was examined by amplifying a DNA fragment spanning the mutation site by reverse transcription followed by PCR (RT-PCR) as described elsewhere (2). Reverse transcription was primed with an oligonucleotide complementary to nt 5360 to 5376 (BW94), and a PCR product was synthesized with oligonucleotides BW35 and BW21 or BW35 and BW94. The PCR product was then cleaved with one of the following pairs of enzymes (numbers in parentheses refer to the position of each restriction site): BamHI(4006)/MscI(4544), MscI(4544)/NcoI(4822), BamHI(4006)/NcoI(4822), or BamHI(4006)/HindIII(5357), depending on the mutant to be analyzed. The appropriate fragments were cloned into a modified pBluescript cut with the same enzymes, and the insert sequences were determined for randomly selected clones with a 373 DNA sequencer (Applied Biosystems).

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Mutations in the conserved portion of the BWYV RTD. Sequence comparisons among members of the Luteoviridae reveal extensive amino acid sequence similarities within the N-terminal half of the RTD (22). Point mutations were generated in this region of the BWYV RTD at or near five positions that were conserved in all sequenced poleroviruses. In selecting the targets, preference was given to amino acids with charged side chains, as we reasoned that such residues are more likely to be exposed on the surface of the protein when it is folded into the native configuration. In the following discussion, amino acid substitutions within the RTD will be referred to by the following convention: XaZ, where X refers to the wild-type residue at position a of the RTD and Z refers to the substituted amino acid.

View larger version (37K): [in this window] [in a new window]   FIG. 2.   Immunodetection of the BWYV RT protein (P74) and major coat protein (P22.5) in transcript-infected C. quinoa protoplasts. Protein extracts of mock-inoculated protoplasts (left-hand lane) or protoplasts inoculated with the indicated BWYV transcript were separated by SDS-PAGE and blotted to nitrocellulose. The nitrocellulose was cut in two; the upper part was probed with an antiserum specific for the RTD, while the lower part was probed with an antiserum specific for P22.5. To the right are indicated the positions of P22.5, P74, and P53, the C-terminally truncated form of the RT protein present in purified virus (loaded in the right-hand lane). Positions of mobility markers (105, 82, 49, 33.5, 28.5, and 19.2 kDa) are shown to the left.

Accumulation of virus in agroinfected plants. Full-length viral cDNAs containing the various RTD mutations were moved into the binary vector pBin19 under control of the cauliflower mosaic virus 35S promoter, and the resulting constructs were introduced into A. tumefaciens. Nicotiana clevelandii plants were inoculated with recombinant agrobacteria harboring either the wild-type cDNA (BW₀) or one of the five above-described mutants. The content of virus antigen was assayed by ELISA on tissue samples from upper noninoculated leaves of the agroinoculated plants (18 to 21 plants tested for each construct) at 5 weeks postinoculation (p.i.).

View larger version (44K): [in this window] [in a new window]   FIG. 3.   Immunodetection of the BWYV RT protein (P74 or P53) and major coat protein (P22.5) in protein extracts from agroinfected N. clevelandii (A) and in purified virus purified from agroinfected N. clevelandii (B). The plants were agroinfected with the indicated constructs, and Western blots were prepared as described in the legend to Fig. 2. In each panel, the upper part of the blot was probed with the RTD-specific antiserum and the lower part was probed with the P22.5-specific serum. The major bands are identified to the right. Mobility markers given to the left in A are as in Fig. 2, and those in panel B correspond to the five smallest markers in Fig. 2.

Sequence of progeny virus in agroinfected plants. The stability of the engineered RTD point mutations during virus multiplication in the agroinfected plants was investigated by RT-PCR. Total RNA was isolated from systemically infected leaves of two plants which had been agroinfected with each mutant, and a region of the genome encompassing the mutation was selectively amplified by RT-PCR. The amplified cDNA (nt 4006 to 4544 for BW5.121, BW5.123, and BW5.125; nt 4006 to 5327 for BW5.127; nt 4545 to 4822 for BW5.129) was then cloned, and inserts from randomly selected clones were sequenced.

View larger version (24K): [in this window] [in a new window]   FIG. 4.   Distribution of nucleotide mutations detected in progeny viral RNA following agroinfection of N. clevelandii with wild-type virus or an RTD mutant (A) and after aphid transmission to M. perfoliata (B). For each mutant, the horizontal line indicates the portion of the RTD sequence which was sequenced, and the number to the right indicates how many RT-PCR clones were analyzed. For mutant BW5.127, the progeny in agroinfected N. clevelandii was analyzed at both 4 and 7 weeks p.i. Positions of the primary mutations in the mutant clones are symbolized by stars. An open star indicates that the primary mutations was conserved in the progeny RT-PCR clones; a filled star indicates that the primary mutations has undergone reversion or pseudoreversion in some or all of the progeny RT-PCR clones. Circles symbolize second-site nucleotide mutations in the progeny RT-PCR clones; open circles denote silent mutations, and filled circles represent mutations which result in an amino acid change. The position of the PVT patch second-site mutations is underlined for mutants BW5.121 and BW5.123.

(i) BW5.121. For BW5.121, the original R24A mutation was maintained in each of the 12 RT-PCR clones characterized. In two clones, a second-site mutation, P32L or T34I, was present near the primary mutation site (Fig. 5A). For brevity, we shall refer to the wild-type sequence PVT(32-34) to which these mutations map as the PVT patch.

View larger version (25K): [in this window] [in a new window]   FIG. 5.   Sequences in the vicinity of the primary mutations of progeny RT-PCR clones following agroinfection (A) or successful aphid transmission (B) with mutants BW5.121 and BW5.123. The wild-type (BW0) RTD sequence is shown at the top; the amino acid(s) targeted in each mutant is indicated in bold. The sequences of RT-PCR progeny obtained from different plants are given below, with the number of times a particular sequence variant was observed indicated to the right. For the transmission experiments, the aphids were rendered viruliferous either by membrane feeding on a purified virus solution or by microinjection of purified virus into the hemocoel. The three amino acids of the PVT patch are identified at the top.

(ii) BW5.123. The primary mutations E59A and D60A were conserved in all 10 RT-PCR clones analyzed, but four of them also contained a second-site mutation, P32L (Fig. 5A). Interestingly, this is the same as one of the PVT patch substitutions observed in BW5.121, although in this case the PVT patch is 26 nt upstream of the primary mutation site at positions 59 and 60.

(iii) BW5.125. The primary mutation K113A/D114A was maintained in all 10 RT-PCR clones analyzed, and no significant second-site mutations were noted (Fig. 4A).

(iv) BW5.127. As shown above, plants agroinfected with BW5.127 initially accumulated low levels of virus, but near-wild-type levels appeared at later times. This observation suggests that the primary K200A/Y201D mutation had affected a motif in the RTD important for virus accumulation but that modified forms of the virus which overcome this defect appear later in infection. To test this hypothesis, viral RT-PCR clones were produced from RNA extracted from leaves taken either early (4 weeks) or later (7 weeks) following agroinfection. The sequence analysis revealed that both of the primary mutations (K200A and Y201D) were present in 20 of 21 RT-PCR clones from three different plants at 4 weeks p.i. (Fig. 6A). In the single exception, D201 had been replaced by N in a clone derived from plant 9. We also detected the mutation S206L in two other clones from plant 9 at 4 weeks p.i. (Fig. 6A), but the significance, if any, of this particular second-site substitution has not been further investigated.

View larger version (21K): [in this window] [in a new window]   FIG. 6.   Sequences in the vicinity of the primary mutations of progeny RT-PCR clones following agroinfection (A) and successful aphid transmission (B) of BW5.127. The wild-type RTD sequence is shown at the top, and the amino acids targeted in the mutant are indicated in bold. The sequences of RT-PCR progeny obtained from different plants are given below, with the number of times a particular sequence variant was observed indicated to the right. Other details are as for Fig. 5.

(v) BW5.129. All of the seven RT-PCR clones examined for BW5.129 retained the primary mutations D225A/E226A and no modifications were noted elsewhere in the region sequenced (Fig. 4A).

Aphid transmission from extracts of virus-infected protoplasts. Extracts of transcript-infected protoplasts were used as a virus source in some transmission experiments. The aphids were allowed a 24-h acquisition access period (AAP) on the protoplast extract. The aphids were then transferred to healthy Montia perfoliata plants for a 4-day inoculation access period (IAP), and the plants were assayed for viral infection by ELISA 3 to 4 weeks later. In such experiments, only mutant BW5.125 was efficiently transmitted (Table 1). Mutant BW5.129 was weakly transmitted, infecting only 3 of 26 test plants following challenge with 30 aphids per test plant. No transmission events were recorded for BW5.121, BW5.123, or BW5.127 even though 100% transmission efficiencies were routinely observed in parallel tests with extracts of BW₀-infected protoplasts (Table 1).

Aphid transmission of virus from agroinfected plants. In other transmission experiments, young, fully expanded leaves of agroinfected N. clevelandii (4 to 6 weeks p.i.) or a solution of purified virus prepared from agroinfected plants was used as a virus source. Nonviruliferous M. persicae nymphs were allowed a 24-h AAP on the leaves or the purified virus solution. Then, 8 or 30 aphids were transferred to healthy M. perfoliata for a 4-day IAP, and the plants were assayed for infection as described above. The tests (Table 1) revealed that for both types of inoculum, mutants BW5.125 and BW5.129 were transmitted with efficiencies similar to that observed in parallel experiments with BW₀. Mutants BW5.121 and BW5.127 were transmitted with intermediate efficiency. Transmission of BW5.123, on the other hand, occurred at a very low rate even though a high inoculation level (30 aphids/plant) was used; only 1 of 8 test plants became infected when BW5.123-agro-infected plants were used as the virus source, and only 2 of 34 plants became infected by aphids which had been given a high concentration (25 µg/ml) of purified virus (Table 1).

Transmission by virus-microinjected aphids. Experiments were carried out to determine if the lower transmissibility observed when the aphids were allowed to acquire BW5.121, BW5.123, and BW5.127 from leaves of agroinfected plants or by membrane feeding on purified virus was due to a defect in a step or steps of the mutant's circuit through the aphid subsequent to passage of the virions from the intestine into the hemocoel. To this end, 0.25 to 0.5 ng of purified wild-type or mutant virus was microinjected into the hemocoel of M. persicae nymphs. The nymphs were then placed on test plants (five aphids per plant) for a 4-day IAP, and infection was scored by ELISA 4 weeks later. BW5.121, BW5.123, and BW5.127 had transmission efficiencies similar to that of wild-type virus (Table 2), suggesting that the RTD mutations in question interfere with the initial step of the transmission circuit, i.e., movement of virions through the intestinal epithelial cell barrier.

Analysis of progeny virus after aphid transmission. The foregoing experiments illustrate that except for BW5.125 (which is readily transmitted from both sources), transmission of the RTD mutants is more efficient following acquisition access to virus from agroinfected plants than it is when the virus is offered as an extract of transcript-infected protoplasts. A plausible explanation for this difference is that the mutant virus is defective in one or more steps of the transmission process but reversions or compensatory mutations which permit transmission have the time to appear during the 4- to 6-week duration of an agroinfection experiment. Furthermore, selective pressure for appearance of compensatory mutations in whole plants could also be exerted by a requirement for a functional RTD for efficient virus movement in planta.

(i) BW5.121. The primary mutation (R24A) was strictly conserved in the aphid-infected plants, but proximal second-site amino acid substitutions were detected in all but one of the 14 RT-PCR clones examined (Fig. 5B). These second-site mutations (P32L in seven clones; T34I in six clones) were the same PVT patch mutations encountered as minor components of the sequence population in the agroinfected plants (Fig. 5A). We conclude that strong selective pressure for appearance of the PVT patch mutations is exerted during aphid transmission of the virus.

(ii) BW5.123. The situation with BW5.123 was more complex. Here, analysis of RT-PCR clones obtained from two plants inoculated with aphids which had been membrane-fed purified virus revealed that one of the primary mutations had undergone a modification: the first A in the mutant sequence AA(59-60) had been replaced by T in plant 2 or had reverted to the wild-type residue E59 in plant 1 (Fig. 5B). To determine if the partial reversion EA(59-60) found in plant 1 restored full transmissibility of the virus, this plant was used as virus source for a second round of aphid transmission. Transmission to eight new M. perfoliata plants occurred with 100% efficiency with both 8 and 30 aphids per test plant. The progeny viral RNA was extracted from two of these plants, and nine RT-PCR clones were sequenced. EA(59-60) was present in all progeny clones, and no mutations elsewhere were noted (data not shown).

(iii) BW5.125. The primary mutation (K113A/D114A) was conserved in each of the 12 clones analyzed, and no second-site amino acid mutations were noted elsewhere in the region sequenced (Fig. 4B). We conclude that the particular motif targeted in BW5.125 is not required for efficient aphid acquisition or transmission.

(iv) BW5.127. For mutant BW5.127 (K200A/Y201D), sequence analysis of progeny RT-PCR products following transmission revealed that the K200A primary mutation was conserved in 14 of the 18 clones. The four exceptions, with a V at position 200, all derived from the same plant (Fig. 6B). We conclude that K200 is dispensable for aphid transmission. At position 201, on the other hand, an amino acid with a phenolic side chain (F or Y) was present in all but one of the progeny clones. The fact that only 1 of 18 RT-PCR clones examined after aphid transmission contained N201 even though this substitution was relatively abundant in BW5.127-agroinfected plants (Fig. 6A) suggests that selective pressure against retention of the N201 variant is applied during transmission.

(v) BW5.129. It was shown above that BW5.129 was transmitted poorly when the virus was supplied to the aphids as an extract of transcript-infected protoplasts but that transmission was efficient when aphids were allowed to acquire virus from agroinfected leaves or by membrane feeding of purified virus (Table 1). We anticipated that the efficient transmission observed in the latter situation would be associated with reversion or compensatory mutations in the BW5.129 genome. However, when the progeny RNA following successful transmission was characterized by RT-PCR, the original D225A/E226A mutations were retained in each of the 11 clones examined (Fig. 4B). Furthermore, although several second-site amino acid substitutions or silent mutations were observed in eight of the progeny RT-PCR clones, none were present more than once except for a silent mutation replacing T(4472) by C, which was detected twice (Fig. 4). Thus, although RTD second-site mutations were relatively frequent in the aphid-transmitted progeny of BW5.129, there was no obvious candidate for a compensatory mutation which could restore efficient transmissibility, as was observed after transmission of BW5.121 and BW5.123.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

In this report we have shown that introduction of point mutations in the conserved portion of the BWYV RTD often leads to the appearance of second-site mutations or pseudoreversions which restore wild-type or near-wild-type function to the virus. This observation suggests that there is considerable structural redundancy in the RTD; that is, the overall folding pattern can tolerate modifications even when highly conserved residues are targeted. It is evident that analysis of such compensatory mutations may help identify motifs in the RTD which are implicated in virus accumulation in planta and in aphid transmission.

The appearance of compensatory mutations in our experiments is certainly related to the relatively high mutation rate characteristic of RNA viruses (8). Even though the inoculum used to agroinfect plants was derived from a cDNA clone and was hence perfectly uniform at the start, sequence variants which are neutral or only slightly deleterious can accumulate during the 4- to 6-week period of virus multiplication in plants. When point mutations are introduced intentionally into the RTD, several scenarios can be envisaged. (i) If the mutation is neutral, the mutant virus will be expected to persist and have properties similar to those of the wild type. This is apparently the case with BW5.125. (ii) If the mutation interferes with viral movement in planta, then the virus will accumulate to low levels following agroinoculation unless reversion or a compensatory mutation restores the movement function. Mutant BW5.127 belongs to this class, at least with respect to the Y201D substitution. (iii) If the mutation does not greatly affect virus accumulation in planta but interferes with aphid transmission of the virus, then transmission efficiency will be reduced and viral genomes carrying reversions or compensatory mutations will appear in the progeny after successful transmission events. BW5.121 and BW5.123 both belong to this category.

At present, BW5.129 cannot be classified. The mutant virus multiplied efficiently in planta following agroinfection. The primary mutations were strictly retained in the progeny virus in the agroinfected plants, and no notable second-site mutations were observed. The primary mutations were also retained following aphid transmission of virus from agroinfected plants, but the high transmission efficiency, compared to the inefficient transmission of virus from infected protoplast extracts, suggested that a compensatory mutation would be present in the progeny. However, although the progeny virus population contained a number of second-site amino acid substitutions and silent mutations in the RTD, there was no obvious pattern in their distribution pointing to a particular second-site mutation as being important. Possibly, there are several distinct second-site mutations in the RTD which can act in concert to compensate for the primary BW5.129 mutations. Alternatively, the compensatory second-site mutations may lie elsewhere in the genome. Further experiments will be necessary to distinguish between these possibilities.

One noteworthy feature of our experiments is that second-site mutations and pseudoreversions were frequently observed but that true reversions were uncommon, being detected only at Y201 for some of the BW5.127 progeny. The low frequency of true revertants is probably related to the fact that the amino acid substitutions introduced into the mutants always involved at least two nucleotide substitutions, generally transversions. The various second-site amino acid substitutions or pseudoreversions which restored function, on the other hand, generally involved a single nucleotide change, and this was almost always a transition. For example, the P32L and T34I second-site mutations in the PVT patch both arose by a T-for-C substitution in the codon. For a number of RNA viruses, it has been demonstrated that transition mutations occur at higher frequency than transversions (17, 19, 28). If the BWYV replicase displays a similar bias in transition/transversion frequency, transitions will be expected to predominate in the pool of potential second-site mutations.

Another unanticipated finding was the ability of the same second-site mutations in the PVT patch (P32I and T34L) to compensate for two distinct primary mutations (R24A in BW5.121 and E59A/D60A in BW5.123) at sites separated by 34 residues. Possibly, R24 and ED(59-60) are brought into proximity during folding of the wild-type RTD, and the PVT patch mutations permit subtle structural adjustments which restore function in the presence of either mutation. It is also possible that the PVT patch mutations act as global stabilizers, i.e., as mutations which augment the stability of the protein (21). If present as a second-site mutation, a global stabilizer may compensate for a primary mutation whose principal effect is to destabilize the protein. It appears unlikely, however, that the PVT patch mutations can function solely as global stabilizers, as it is difficult to imagine how a global stabilizer could act in a differential manner at different steps of the acquisition/transmission process, as observed for BW5.123-P32L. Evidently, although identification of second-site compensatory changes can provide preliminary information about structure-function relationships, knowledge of the three-dimensional structure of the RTD will be essential for an in-depth interpretation of the data.

Finally, the double mutant BW5.123-P32L, which is apparently unable to cross from the intestine into the hemocoel, could provide a useful probe for dissecting the acquisition process. Acquisition of BWYV involves receptor-mediated endocytosis of virus particles at the apical plasmalemma of intestinal epithelial cells, unidirectional intracellular transport through these cells in vesicles, and exocytosis into the hemocoel (10, 11). Electron microscopic observations of thin sections of intestinal epithelia of M. persicae which have been membrane fed with this mutant may tell us at what stage of this process the virus is blocked and so provide information about receptor localization.