Movement Protein of a Closterovirus Is a Type III Integral Transmembrane Protein Localized to the Endoplasmic Reticulum

Cell-to-cell movement of beet yellows closterovirus requiresfour structural proteins and a 6-kDa protein (p6) that is aconventional, nonstructural movement protein. Here we demonstratethat either virus infection or p6 overexpression results inassociation of p6 with the rough endoplasmic reticulum. Thep6 protein possesses a single-span, transmembrane, N-terminaldomain and a hydrophilic, C-terminal domain that is localizedon the cytoplasmic face of the endoplasmic reticulum. In theinfected cells, p6 forms a disulfide bridge via a cysteine residuelocated near the protein's N terminus. Mutagenic analyses indicatedthat each of the p6 domains, as well as protein dimerization, isessential for p6 function in virus movement.

INTRODUCTION

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Introduction

Transport of plant viruses within and between cells is an activeprocess that requires the function of virus-coded movement proteins(MPs). By definition, MPs are specialized proteins that areessential for the translocation of viral genomes or virions,but they are not required for virus genome replication or encapsidation.Viral MPs belong to several distinct protein families, eachof which seems to exhibit a unique functional profile (10, 26).Many virus genera possess not one but two or three MPs. In addition,cell-to-cell movement of some viruses requires proteins whoseprimary functions are in genome replication or encapsidation (8, 9, 21).

Among several present models of virus movement, two have approachedcanonic status (10, 26). One is a Tobacco mosaic virus (TMV)model (4). The only TMV MP, the 30-kDa protein p30, is ableto bind viral RNA and guide it through the plasmodesmata (13).Its additional activities include modification of plasmodesmatalgating properties and interactions with microtubules, actinmicrofilaments, endoplasmic reticulum (ER) (28, 31, 38), anda cell wall-specific host enzyme (12).However, the exact mechanisticcontributions of these MP associations to intracellular movementare a matter of debate (6, 17, 41). Likewise, the transportmechanism of the RNA-MP complex through plasmodesmata largely remainsa mystery. The leading model proposes that TMV-type MPs recruit apreexisting host machinery for intercellular trafficking (19, 27).Interestingly, both rod-shaped RNA viruses related to TMV andseveral icosahedral RNA and single-stranded DNA viruses appearto follow this movement paradigm (16, 26).

The second well-recognized model applies to several familiesof the icosahedral RNA viruses and pararetroviruses (35, 42).The MPs of these viruses reorganize plasmodesmata by inducingformation of the tubules through which mature virions translocatefrom cell to cell. The MP and endomembrane secretion systemappear to be sufficient for tubule formation, whereas intactcytoskeleton is required for proper positioning of the tubulesrelative to plasmodesmata (24).

Mounting evidence indicates that the filamentous potexvirusesdo not fit in any of the abovementioned models. The 25-kDa MP(p25) of Potato virus X (PVX) possesses nucleoside triphosphataseand RNA helicase activities and is able to disassemble virionsin a polar manner (29). p25 was the first viral MP for whicha role in suppression of the host RNA silencing defense responsewas demonstrated (44). In addition to p25, the quadripartitePVX movement machinery includes two membrane-bound MPs and acapsid protein (CP), each of which is essential, but not sufficient,for virus translocation (11, 23).

The family Closteroviridae in general and the Beet yellows virus (BYV)in particular occupy a special niche among models of plant virologydue to their large RNA genomes, exceptionally long filamentous virions,and a five-component machinery for cell-to-cell movement (14).Four of the BYV movement-associated proteins are the virioncomponents. One is a major CP which encapsidates most of thevirion RNA. The three others are the minor CP (CPm), a 64-kDaprotein (p64), and a homolog of the $~$ 70-kDa heat shock proteins(Hsp70h). Remarkably, CPm, p64, and Hsp70h assemble virion tailsthat were proposed to function as a specialized movement device (3, 30).The only "conventional" BYV MP is a 6-kDa hydrophobic protein (p6).Although p6 is not required for assembly of the movement-competent,tailed virions, it is essential for BYV movement from cell tocell (2, 3, 33).

In this work, we demonstrate that BYV p6 is inserted into ERmembranes with its C-terminal hydrophilic domain facing thecytosol. The Cys-3 residue of p6 is present within the ER lumenand is involved in the formation of the disulfide bond. Mutationalanalysis of p6 revealed that the short luminal, transmembrane,and cytosolic regions of this protein are each essential forp6 function in BYV cell-to-cell movement.

MATERIALS AND METHODS

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Materials and Methods

Isolation and chemical treatments of the microsomal fractions. Symptomatic leaves from BYV-infected Nicotiana benthamiana plantswere ground in lysis buffer (20 mM HEPES [pH 6.8], 150 mM potassiumacetate, 250 mM mannitol, 1 mM MgCl₂), and the homogenate wasclarified by centrifugation at 3,000 x g for 10 min at 4°C.The supernatant was centrifuged at 30,000 x g for 1 h at 4°Cto yield the soluble (S30) and the crude (P30) microsomal fractions.Microsomal pellets were resuspended in 10 vol of 100 mM Na₂CO₃(pH 11), 4 M urea, or original lysis buffer and incubated for30 min on ice (39).

Membranes were collected by centrifugation at 30,000 x g for30 min and resuspended in the original volume of lysis buffer.The S30 and P30 fractions were prepared and boiled in sodiumdodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)sample buffer with or without dithiothreitol (DTT) as indicatedin Results.

Alternatively, P30 was resuspended in lysis buffer containing1% Triton X-100, incubated on ice for 30 min, and centrifugedat 30,000 x g for 1 h. The resulting pellet was resuspendedin lysis buffer in a volume equal to that of the original sample.

Treatment with Triton X-114 was performed by resuspending P30in lysis buffer containing 1% Triton X-114, clarifying by centrifugationat 0°C, and incubating the lysate at 37°C for 10 minwith subsequent centrifugation at 10,000 x g at room temperatureto allow separation of aqueous and organic phases (5, 39). Thelower phase, corresponding to the detergent-rich fraction, waswashed by addition of fresh buffer lacking Triton X-114 andvortexing. The tube was placed on ice for 10 min, and phaseseparation was repeated.

Sucrose gradient fractionation, immunoblotting, and proteinase treatments. Plant material was harvested and ground in lysis buffer containingeither 0.1 or 5 mM MgCl₂. P30 prepared as described above wasloaded on top of 20 to 60% linear sucrose gradients containinglysis buffer with corresponding concentrations of MgCl₂ (39, 46).Gradients were centrifuged for 16 h at 100,000 x g in a BeckmanSW40 rotor at 4°C, and 15 fractions were collected startingfrom the top. Aliquots from each fraction were separated using SDS-PAGE.Immunoblot analyses were done using rabbit polyclonal antibodiesto BYV p6 (33), BiP (a gift from Maarten Chrispeels, Universityof California, San Diego), or green fluorescent protein (GFP)(Living Color antibodies; Clontech, San Jose, Calif.).

For proteinase K treatments, microsomal pellets were resuspendedin lysis buffer and loaded on top of discontinuous sucrose gradientsconsisting of 20 and 60% sucrose. Gradients were centrifugedfor 2 h at 100,000 x g. A fraction containing closed-cell vesicles originatingfrom disrupted ER (top of 60% sucrose phase) was collected.Aliquots of this fraction were treated with 100 µg ofproteinase K/ml at 0°C in the presence or absence of 1%Triton X-100 (37). After 20 min, the reactions were quenchedby the addition of 2 mM phenylmethylsulfonyl fluoride and theproducts were used for immunoblotting.

Molecular cloning and analyses of the mutant viruses and p6 variants. The alanine-scanning and premature stop codon mutations listedin Table 1 were introduced into the p6 open reading frame (ORF)by using p65 M plasmid as described previously (2). The SnaBI-NdeIfragments from the resulting mutant plasmids were engineeredto the pBYV-GFP plasmid that harbored a full-length cDNA cloneof BYV tagged by the insertion of the GFP expression cassette(34). The corresponding in vitro RNA transcripts were used toinoculate Claytonia perfoliata plants, and the cell-to-cellmovement of the resulting viruses was assayed as described previously (34).

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TABLE 1. Mutation analysis of p6 function in BYV cell-to-cell movement

Alternatively, wild-type or mutant p6 ORFs were inserted intoa pCB-derived minibinary vector for Agrobacterium-mediated overexpressionin the N. benthamiana leaves (32, 36). To generate fusion ofthe p6 and GFP ORFs (p6/GFP), enhanced GFP ORF(Clontech) wasinserted in-frame downstream from the coding sequence of p6.Analogously, enhanced GFP ORF was inserted in-frame upstreamfrom the sequence encoding the C-terminal, hydrophilic domainof p6 (codons 32 to 54) to generate a fusion ORF, designatedGFP/C-p6.

To transiently express p6 variants, the minibinary plasmidswere introduced into Agrobacterium tumefaciens strain C58 GV2260by electroporation. Leaves of N. benthamiana plants were infiltratedwith the resulting agrobacteria and used either for immunoblotanalyses or for microscopic detection of the p6/GFP fusion products(32). Confocal laser scanning microscopy was done 2 days afterleaf infiltration by using an inverted Leica DMIRBE microscopeequipped with a TCS4D laser and a band-pass fluorescein isothiocyanate filter.

RESULTS

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Results

BYV p6 is an integral membrane protein. Computer analysis of the p6 amino acid sequence consistentlypredicted that p6 is a membrane protein with a single, ${alpha}$ -helicaltransmembrane domain spanning residues 8 to 32 (Dense AlignmentSurface [DAS] transmembrane prediction server), 9 to 32 (Kyte-Doolittle hydrophobicityplot), or 12 to 31 (PHD program). Subcellular fractionationof the extracts derived from the BYV-infected plant tissue wasused to test computer predictions. The immunoblot analysis revealedthat virtually all p6 was present in a pellet fraction (P30)following centrifugation at 30,000 x g (Fig. 1A). Because thisfraction contains membrane-derived microsomes, we assumed thatp6 is indeed a membrane-associated protein. This assumptionwas supported by solubilization of p6 by the treatment of P30with the nonionic detergent Triton X-100 (Fig. 1A) (37). Todistinguish between the luminal and membrane-associated localizationsof p6, P30 was treated with Na₂CO₃ (pH 11). Such treatment rendersmicrosomes to open membranous sheets, thus releasing the solubleluminal proteins (39). Because p6 remained in the pellet (Fig.1B), we concluded that it is tightly associated with the membranes.To determine whether p6 is a peripheral membrane protein, theP30 fraction was treated with urea (37, 39). This treatmentdid not release p6 (Fig. 1B), suggesting that p6 is anchoredwithin the membrane. To confirm this conclusion, P30 was alsotreated with Triton X-114, a detergent that forms a separatephase to which the membrane lipids and hydrophobic proteinsare partitioned (5). Because p6 was detected in the hydrophobic,but not the aqueous, fraction (Fig. 1C), we concluded that p6 isan integral membrane protein.

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FIG. 1. Immunoblot analyses, using anti-p6 serum, of the extracts from BYV-infected plants. Lanes: Total, protein extract prior to fractionation; 3K, supernatant following extract centrifugation at 3,000 x g; S and P, supernatant and pellet, respectively, following centrifugation at 30,000 x g; AP and OP, aqueous and organic phases, respectively, following extraction with Triton X-114. Samples treated with Triton X-100, urea, or Na₂CO₃ buffer are marked accordingly.

p6 resides in the rough ER. Sucrose density gradient centrifugation in the presence of 5mM MgCl₂ was employed to fractionate p6-containing membranesderived from the BYV-infected plants. Immunoblot analysis ofthe gradient fractions using p6-specific antibody revealed thepeak of p6 in fractions 9 to 11 (Fig. 2, top), a pattern suggestiveof p6 association with the ER membranes. Indeed, probing thesame gradient fractions with the antibody to a residential ERprotein, BiP, demonstrated cofractionation of BiP and p6 (Fig.2, middle). Furthermore, in the presence of 0.1 mM MgCl₂, bothp6 and BiP peaked in fractions 7 to 9 (Fig. 2). Similar resultswere obtained with the ER-targeted GFP (Fig. 2, bottom) (18).Because such mobility shift due to release of the ER-associatedribosomes is characteristic of the rough ER (37, 39, 46), weconcluded that p6 is likely present in the rough ER. It is worthnoting that both BiP and ER-targeted GFP, but not p6, were alsoobserved at the top of the gradient (Fig. 2). This could bedue to release of ER-luminal BiP and GFP from the popped ER vesicles,whereas no such release could be expected for p6 if it were anintegral membrane protein.

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FIG. 2. Immunoblot analyses of the protein extracts following separation in the sucrose gradients, with the fraction numbers shown at the top. The types of antisera used for analysis are indicated at left, and MgCl₂ concentrations are shown at right. BiP, ER-resident marker protein.

To visualize p6 within the living cells, we fused GFP to theC terminus of p6 and overexpressed the resulting p6/GFP productin leaf tissue. Confocal laser scanning microscopy revealedthat p6/GFP was localized primarily to a reticulate network,which was indistinguishable from that observed in transgenic 16cplants engineered to express the ER-targeted GFP (Fig. 3A and B).Interestingly, fusion of the C-terminal, hydrophilic domainof p6 to the GFP C terminus (GFP/C-p6) resulted in the uniformdistribution of this product in cytoplasm (Fig. 3C). This resultis compatible with the notion that p6 is targeted to the ERvia its N-terminal, hydrophobic domain rather than via interactionof the C-terminal, hydrophilic domain with the peripheral ERprotein. Taken together, the results presented above establishp6 as an integral membrane protein that is targeted to the roughER in the BYV-infected cells or upon transient expression.

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FIG. 3. (A to C) Confocal laser scanning microscopy analysis of the 16c transgenic plants that express ER-targeted GFP (A), the p6/GFP fusion (B), or the GFP fusion to the C-terminal, hydrophilic domain of p6 (C). The green corresponds to the GFP fluorescence, and the occasional red spots represent the autofluorescent chloroplasts. (D) Amino acid sequence (top) and membrane topology (bottom) of BYV p6. A1 to A12 and arrows indicate the alanine-scanning mutations introduced into indicated positions of p6. Red hexagons indicate premature stop codon mutations replacing the residues shown above.

Membrane topology of p6. A single topogenic domain of p6 functions as both signal andanchor for insertion into membrane. Therefore, p6 could be eithera type II membrane protein with its C terminus located withinthe ER lumen or a type III membrane protein with a luminal Nterminus (40). Whatever is the case, the luminal segment ofp6 must be exposed to an oxidizing environment that promotesdisulfide bond formation via available Cys residues (15). Becausep6 possesses a Cys residue at the third position from the Nterminus (Fig. 3D), whereas the other two cysteines are buriedwithin the membrane, the ability of p6 to dimerize would suggestthat it is a type III membrane protein. To investigate the possibilityof p6 dimerization, we performed an electrophoretic analysisof the protein extracts from BYV-infected plants (Fig. 4A, lanes1 and 2) or plants that transiently expressed p6 (Fig. 4A, lanes9 and 10) under nonreducing versus reducing conditions. Strikingly,for both extracts, these analyses revealed dramatically differentp6 mobilities depending on the presence of DTT in the proteindissociation buffer. Under reducing conditions, p6 migratedas an $~$ 12-kDa protein, whereas its mobility under nonreducingconditions corresponded to an $~$ 18-kDa protein. This result iscompatible with the ability of p6 to dimerize. We assume thatanomalously slow migration of the p6 monomers and dimers isdue to p6 hydrophobicity.

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FIG. 4. (A) Analysis of dimerization of wild-type p6 and three alanine mutants targeting each of the cysteine residues present in p6 (see also Fig. 3D and Table 1). Lanes: p6, agrobacterium-mediated expression of p6; BYV, virus-infected plants. The presence or absence of DTT in the protein dissociation buffer is indicated above the lanes with a plus or minus sign, respectively. D, p6 dimer; M, p6 monomer. Positions of the protein markers are shown at left. In both panels A and B, p6 was detected using immunoblotting and p6 antiserum raised against the C-terminal hydrophilic domain of p6. (B) Dimerization of the p6/GFP fusion product. The designations are the same as in panel A, except for M* and D*, which correspond to a monomer and a dimer formed by the p6/GFP fusion product, respectively. (C) Treatment of the resuspended P30 fraction of the p6-containing protein extracts with proteinase K (PrK) in the presence or absence of Triton X-100 as indicated at the top.

To determine whether, as expected, Cys-3 is required for p6dimerization, we replaced this residue with Ala and expressedthe resulting mutant protein in plants. For comparison, Alawas also substituted for Cys-18 and Cys-23, each of which islocated within the transmembrane domain (Fig. 3D) and not expected toform disulfide bridges. In complete agreement with these predictions,mutation of Cys-3, but not of Cys-18 or Cys-23, resulted inloss of p6 dimerization (Fig. 4A, lanes 3 to 8). Because formationof disulfide bonds catalyzed by protein disulfide isomeraseis feasible only within the ER lumen (15), we concluded that theN-terminal segment of p6 including Cys-3 is indeed present inthis compartment and that p6 is likely a type III membrane protein.

The $~$ 18-kDa p6 dimer could be either a homodimer or a heterodimerformed by p6 with an apparent molecular mass of 12 kDa and byanother protein of $~$ 6 kDa. To distinguish between homo- and heterodimerizationof p6, we compared dimerization of p6 to that of the p6/GFPfusion product (Fig. 4B). The p6/GFP molecular mass of $~$ 38 kDaestimated by SDS-PAGE under reducing conditions (Fig. 4B, lane4) was reasonably close to the sum of the molecular mass ofGFP ( $~$ 27 kDa) and the apparent molecular mass of the p6 monomer( $~$ 12 kDa). If this protein were to form a homodimer, the expectedmolecular mass should be $~$ 76 kDa. Alternatively, if p6/GFP wereto form a heterodimer with the same hypothetical protein asin the case of p6, its estimated molecular mass should be $~$ 44kDa [molecular mass of p6/GFP plus the difference between themolecular masses of the p6 dimer and monomer: 38 kDa + (18 kDa- 12 kDa) = 44 kDa]. Because the estimated molecular mass ofthe p6/GFP product analyzed under nonreducing conditions was $~$ 76 kDa (Fig. 4B, lane 4), it could be assumed that p6/GFP and,likely wild-type p6, form homodimers.

To confirm a type III membrane topology of p6, a sucrose gradient-purifiedmicrosomal fraction derived from plants transiently expressingp6 was treated with proteinase K in the absence or presenceof Triton X-100 (37). It was expected that if the hydrophilicC-terminal domain of p6 is present in the cytoplasm, it wouldbe proteolytically degraded with or without membrane solubilizationby Triton X-100. Conversely, if the C-terminal domain is presentin the ER lumen, it would be protected from digestion in theabsence, but not in the presence, of a detergent. Because the antibodyused to detect p6 was raised against synthetic peptide correspondingto the C-terminal 23 residues of p6 (33), digestion of the C-terminaldomain should abolish p6 immunogenicity. As shown in Fig. 4C,the obtained results are clearly compatible with the former,but not the latter, scenario. That is, p6 lost its C-terminalimmunogenic determinants following proteinase K treatment inboth the absence and presence of the detergent. The same resultswere obtained when trypsin was used for the treatment (datanot shown). Taken together with the ability of p6 to dimerizevia the Cys-3 residue, these data establish p6 as a type III membraneprotein with its hydrophilic, C-terminal region facing the cytosol.

Mutational analysis of p6 function in BYV cell-to-cell movement. Alanine-scanning mutagenesis was used to map structure-to-functionrelations within the p6 molecule. Twelve Ala replacement mutationswere introduced along the entire p6 sequence (Fig. 3D and Table 1).The mutant p6 variants were engineered into a cDNA clone ofBYV tagged via insertion of the GFP reporter to visualize replicationand cell-to-cell movement of the virus in indicator plants (34).Approximately 60 to $~$ 250 of the individual infection foci wereanalyzed for each of the BYV variants.

Each of the three mutations introduced into the short, N-terminal,luminal segment of p6 (A1 to A3) resulted in the dramatic reductionor complete loss of virus ability to move from cell to cell(Table 1). Likewise, all four mutations that targeted the transmembranedomain (A4 to A7) also affected virus movement. Finally, fiveAla replacements of the positively or negatively charged residuesin the C-terminal hydrophilic domain of p6 (A8 to A12) all resultedin reduced movement. These data indicate that p6 function isunusually prone to structural changes within each of the threetopological regions of this membrane protein. It should be emphasizedthat the A2 mutation that replaced a Cys-3 residue involvedin p6 dimerization stands alone in that all of the detected93 infection foci were unicellular. In contrast, each of theother alanine-scanning mutants was able to generate at leasta few multicellular foci (Table 1). This result underscoresthe critical nature of Cys-3 and suggests that dimerization isessential to p6 function in BYV cell-to-cell movement.

To further probe the cytosolic domain of p6, six premature stopcodons were introduced along its length (Fig. 3D). As shownin Table 1, progressive truncation of p6 resulted in gradualloss of its function. Only two mutants that truncated p6, bythree and six residues, were able to move, albeit inefficiently.Even though the length and amino acid sequence of the C-terminaldomain are not conserved among p6 orthologs encoded by the membersof the genus Closterovirus (data not shown), both alanine-scanningand truncation analyses point to very rigid structure-to-functionrelations within this domain.

DISCUSSION

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Discussion

The endomembranes in general and ER in particular play conspicuousyet poorly understood roles in virus-cell interactions. Many,but not all, positive-strand RNA viruses recruit ER for theformation of the membrane-enveloped spherules to which replicationcomplexes are sequestered (1). In addition, several plant virusMPs that function in a TMV-like, comovirus-like, or PVX-likemanner were found in tight association with the ER (4, 22, 23, 25, 29, 45).In most of these cases, the mechanistic significance of theMP-ER association is unclear. The apparent connection of theplasmodesmata with ER elements and the presence of the appressedER in the desmotubule prompted the suggestion that viruses needto modify and/or recruit plasmodesma-associated ER for theirmovement. It has also been demonstrated that pharmacologicalinactivation of membrane trafficking from the ER to the Golgiapparatus and beyond affects localization of the TMV MP to thecell periphery (20). Analogously, it has been demonstrated thatmembrane trafficking is required either for proper targetingof the comoviral MPs or for the ability of MPs to form movement-associatedtubules (24, 35).

The present model of the ER-associated form of the 30-kDa TMVMP features two transmembrane domains, with the termini of bothproteins facing the cytosol (7). A similar model was developedfor the unrelated, 9-kDa MP of a carmovirus (43). So far, the6-kDa BYV MP is unique among other MPs due to its small sizeand a single-span, type III membrane topology (Fig. 3D). However,because p6 forms disulfide bonds via its Cys-3 residue, theresulting homodimer also contains two cytosolic domains.

How does such a small protein molecule provide a critical contributionto the movement of unusually large BYV virions? Although themechanism of p6 action remains a matter of speculation, twopossibilities are compatible with the existing data. The firstassumes that the modification of the ER membranes by p6 is requiredto promote BYV movement. Such a modification could be neededto release nascent virions from ER-derived vesicular aggregates,where BYV RNA is synthesized and likely assembled. p6 couldalso promote virion transport to the cell periphery in associationwith the ER-derived vesicles. A second possibility is that p6modifies the secretion pattern of the infected cell. Such a modificationcould be negative, i.e., suppression of the secretion of theER-synthesized antiviral factors. It also could be positive,i.e., stimulation of the secretion of the factors required forvirus transport. Interestingly, each of the 18 point mutationsintroduced into ER-luminal, transmembrane, or cytosolic segmentsof p6 resulted in partial or complete loss of function. Theseunusually stringent structural requirements for this minuteMP suggest its involvement in critical interactions with thehost and/or viral factors.

Is p6 distributed uniformly within the ER network of the infectedcell, or is it confined to a subset of ER elements? Is p6 capableof association with the Golgi apparatus or plasma membranes?These and other related questions will be addressed by usingp6 fused to a monomeric red fluorescent protein in combinationwith GFP-tagged organelles. In addition, p6 provides an excellentmodel for probing the ER structure and function. In particular,the small size and simple membrane topology of p6 are usefulfor probing the mechanisms of targeting and retention of transmembraneproteins in the ER as well as the structural requirements forproper membrane orientation and disulfide bond formation.

ACKNOWLEDGMENTS

We thank Jim Carrington and Christophe Ritzenthaler for criticalreading of the manuscript, David Baulcombe for 16c transgenicplants, Maarten Chrispeels for the BiP-specific antibody, andJim Haseloff for the ER-targeted GFP.

The research was supported by grants from the National Institutesof Health (R1GM53190) and the U.S. Department of Agriculture(CSREES 2001-35319-10875) to V.V.D.

FOOTNOTES

* Corresponding author. Mailing address: Department of Botany and Plant Pathology, Oregon State University, Cordley Hall 2082, Corvallis, OR 97331. Phone: (541) 737-5472. Fax: (541) 737-3573. E-mail: doljav{at}science.oregonstate.edu.

REFERENCES

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