JOURNAL

OF

Vol. 66, No. 1

VIROLOGY, Jan. 1992, p. 258-269

0022-538X/92/010258-12$02.00/0 Copyright © 1992, American Society for Microbiology

Deletion of the VP16 Open Reading Frame of Herpes Simplex Virus Type 1 STEVEN P. WEINHEIMER,l* BRANIN A. BOYD,' STEPHEN K. DURHAM,2 JAMES L. RESNICK,'t AND DONALD R. O'BOYLE II' Department of Virology' and Department of Experimental Pathology and Electron Microscopy,2 Bristol-Myers Squibb Pharmaceutical Research Institute, P.O. Box 4000, Princeton, New Jersey 085434000 Received 13 June 1991/Accepted 5 October 1991

VP16 (also called Vmw65 and caTIF) is a structural protein of herpes simplex virus type 1 (HSV-1) that trans-induces HSV-1 immediate-early gene transcription. This report describes an HSV-1 VP16 deletion mutant that was constructed and propagated in a cell line transformed with a VP16 expression vector. The VP16 deletion mutant replicated like wild-type HSV-1 during infection of the VP16-expressing cell line. Deletion mutant virions propagated in this cell line contained wild-type, cell-derived VP16 protein that was recruited during virion assembly and was functional for immediate-early gene trans-induction. The mutant failed to replicate during subsequent infection of cells that do not express VP16, as determined in plaque assays and single-step replication assays. The deletion mutant induced nearly normal levels of viral DNA synthesis and capsid production during these infections, but it induced slightly lower levels of viral DNA encapsidation and appeared by transmission electron microscopy to be defective in further steps of virion maturation. A genetic revertant of the deletion mutant that was restored for VP16-coding sequences exhibited fully wild-type replication properties in both VP16-expressing and nonexpressing cells. The absence of VP16 protein synthesis at late times of HSV-1 infection prevents the production of infectious progeny virus and correlates with a profound defect in HSV-1 particle assembly.

Proteins encoded by herpes simplex virus type 1 (HSV-1) are synthesized during lytic infection in a progressive cascade, from immediate-early (IE) to delayed-early (DE) to late viral proteins (17). IE proteins regulate and are required for DE gene expression. DE proteins include enzymes involved in nucleotide metabolism and viral DNA synthesis. Viral DNA replication induces high levels of HSV-1 late gene expression, and late proteins comprise mostly virion structural proteins (reviewed in reference 42). VP16 (also called Vmw65 and a-TIF) is a 65-kDa HSV-1 virion phosphoprotein (23, 25) synthesized during the late phase of HSV-1 gene expression (16). During virion assembly, VP16 is incorporated into the tegument between the capsid and the virion envelope (4, 27). This virion-associated VP16 is subsequently released during infection, whereupon it specifically trans-induces the transcription of viral IE genes (4, 5, 31). The specific induction of IE genes by VP16 is mediated via two cis-acting DNA sequence elements, the TAATGA RAT (R = purine) (24, 41) and GCGGAA (41) consensus sequences of IE promoters. This induction requires the physical interaction of VP16 with cellular factors (11, 19-21, 28, 33, 44), one of which (Oct-1) binds directly to the TAATGARAT sequence element (11, 40). Investigations using HSV mutants encoding altered VP16 proteins have begun to illustrate the extent to which, and in what capacity, VP16 is required for HSV lytic replication. VP16 is encoded by the 490-amino-acid open reading frame (ORF) UL48 of HSV-1 (7, 26, 29). Preston et al. created a viral mutant, in1814, containing a four-amino-acid insertion at position 379 of UL48 (2) that eliminates the transactiva-

tion of IE promoters by VP16 (1, 2). Viral gene expression and viral replication are both greatly reduced relative to what is found with wild-type controls when cells are infected with low numbers of in1814 virions (e.g., 75% of the capsid was filled with electron-dense material. RESULTS Construction of a VP16-expressing cell line. We anticipated that an HSV-1 mutant lacking VP16-coding sequences might grow poorly, if at all, on normal tissue culture host cells. A recombinant cell line was created to propagate such a mutant by transforming Vero cells with a VP16 expression vector, pVP16-9. The insert of pVP16-9 is the expression cassette from pMSVP16, which spans sequences from 76 bp upstream to 118 bp downstream of the VP16 ORF of HSV-1 strain KOS (41). In this cassette, VP16 expression is directed from the Moloney murine sarcoma virus long terminal repeat and terminated by the polyadenylation signals of the HSV-1 thymidine kinase gene (41). VP16 is the only HSV-1 protein known to be encoded by the sequences of pVP16-9 (5, 7, 16, 26, 29). pVP16-9 was cotransfected into Vero cells with pSV2neo (38), which encodes the aminoglycoside phosphotransferase gene, and 30 G418-resistant colonies were isolated (see Materials and Methods). The colonies were screened by Southern blotting for the presence of stably inherited copies of the VP16 expression insert (data not shown). Five transformants were selected from those that tested positive for inheritance of the VP16 expression cassette (-50%). All five were infected with the HSV-2 mutant ts2203 at 34 and 38.6°C to test for trans-complementation of the conditional-lethal defect of this mutant. ts2203 formed plaques at 38.6°C in three of the five cell lines tested (data not shown). One transformant, designated Vero-16-8 (16-8), supported the same level of ts2203 plaque formation at the nonpermissive (38.6°C) and permissive (34°C) temperatures. We concluded that this complementation was due to the presence of VP16 protein provided in trans and it did not result from marker rescue by the resident cellular copies of the VP16 gene fragment, since wild-type rescuants could not be detected among progeny virus produced in virus yield assays (data not shown). These results suggest that the ts2203 mutation alters the activity of a VP16 homolog of HSV-2 and that 16-8 cells express functional VP16 capable of rescuing the growth of a VP16-defective strain of HSV. A deletion/substitution allele of VP16. We employed standard techniques for HSV-1 mutagenesis to create a VP16 deletion mutant by site-specific recombination (13, 35). The first necessary step was to construct a plasmid-encoded copy of the mutation to be introduced into the HSV-1 chromosome. The parental plasmid for mutagenesis was a subclone of the EcoRI I fragment of HSV-1 strain KOS (Fig. 1, top panel). This subclone, designated pVP16-KOS, spans from an XhoI site 455 bp 5' of the VP16 ATG initiation codon to a PstI site 944 bp 3' of the VP16 TAG termination codon (Fig. 1, second panel). VP16-coding sequences in pVP16KOS were deleted by digestion with EcoRV and StyI, and a synthetic BglII linker was inserted (Fig. 1, third panel). This removed sequences from 28 bp 5' of the VP16 initiation codon to 20 bp 5' of the VP16 termination codon, eliminating

VOL. 66, 1992

VP16 DELETION MUTANT OF HSV-1

pSG22 HSV-1 MAP UNITS

v

pBR32S

ORF .632

.720

SaclI Styl

EcoRV Ap9I

pVP16-KOS

ATG

PstI

pTZ18R

VP16 ORF

7~~~Bg

\ XbaI pVP16RVA Sty

261

\

\1/ /

/18R

EcoRI

\PstI

EcoRI Xba1 pTZ/18R / CK -GALACTosiDASE _ pVP16Af Gal FIG. 1. Construction of pVP16APgal, a plasmid-encoded VP16 deletion/substitution mutant. Plasmid designations are shown at left. Vectors (solid lines) are indicated at right. Boxed segments represent plasmid inserts. Location of the VP16 ORF is indicated, as are the positions of VP16 ATG and TAG codons. Hatched boxes indicate HSV-1 sequences flanking the VP16 ORF that were retained during subcloning and mutagenesis. Top panel: pSG22 (12), the EcoRI fragment I of HSV-1 strain KOS (map units 0.632 to 0.720) containing the VP16 ORF. Second panel: pVP16-KOS, a 2.8-kb XhoI-PstI fragment of pSG22 subcloned in pTZ18R. The XhoI site was eliminated by ligation to the Sall site of the polylinker. Third panel: pVP16RVASty, a deletion of the 1.5-kb EcoRV-StyI fragment of pVP16-KOS (indicated by dashed lines). A BglII linker was inserted in place of the deletion. Bottom panel: pVP16A13gal, an insertion of the 4.3-kb BamHI fragment of pD6P at the BglII site of pVP16RVASty (also indicated by dashed lines). The orientation of P-galactosidase transcription is indicated with a bent arrow.

the initiation codon and all but the 8 most carboxyl terminal amino acid codons of the VP16 ORF. The resulting construct, pVP16RVASty, retained 428 bp of flanking HSV-1 sequences 5' of the deletion and 975 bp of flanking HSV-1 sequences 3' of the deletion. A DNA cassette containing the HSV-1 ICP6 promoter driving expression of an mRNA encoding the bacterial enzyme ,B-galactosidase (13) was inserted at the BglII site of pVP16RVASty (Fig. 1, bottom panel). This final construct, designated pVP16APgal, was used for mutagenesis of the HSV-1 chromosome. According to the DNA sequence analyses of McGeoch and colleagues (26), the VP16 ORF is flanked on either side by an ORF, UL49 in the 5' direction and UL47 in the 3' direction, and both lie in the same orientation as the VP16 ORF. The 5' boundary of the VP16 deletion lies 380 bp downstream of the predicted carboxyl terminus of UL49 and 355 bp downstream of the putative polyadenylation signal for UL49 mRNA. The 3' boundary of the VP16 deletion lies 319 bp upstream of the mapped 5' end of UL47 mRNA and 516 bp upstream of the predicted amino terminus of UL47 (26). Precise deletion of the VP16-coding sequences in pVP16APgal was carried out to avoid causing secondary defects in UL47 or UL49 gene expression during HSV-1 infection.

Isolation of a recombinant HSV-1 VP16 deletion/substitution mutant. The flanking HSV-1 sequences retained 5' and 3' of the VP16 deletion/substitution mutation in pVP16APgal were used to drive homologous recombination at the chromosomal VP16 locus of HSV-1. Infectious HSV-1 virion DNA (strain KOS) was cotransfected into 16-8 cells with the plasmid insert of pVP16APgal. The virus progeny from this transfection were used to inoculate additional 16-8 monolayers, and the resulting plaques were stained with X-Gal, a chromogenic substrate of ,B-galactosidase. Ten positively stained plaques derived from two independent transfections were selected, and the recombinant virus isolates were plaque purified (see Materials and Methods). Five of these isolates were analyzed further by Southern blotting (37), as described below. DNA from partially purified virions was digested with restriction enzyme BamHI or EcoRI (see Materials and Methods), and three identical Southern blot filters were prepared from 0.8% agarose gels (37). The hybridization results for all five mutant virus isolates were identical (data not shown); the results observed for one isolate, designated 8MA, are presented in Fig. 2A. One of the three filters was probed with an ApaI-SacII fragment of pVP16-KOS that lies internal to the VP16 deletion boundaries to detect VP16-

262

WEINHEIMER ET AL.

A.

._-

J. VIROL.

B.

8MA sS

CODING SEQUENCE

pN T1 6-K()S SAC 11

.-x1PA. I

L N-TI 6 O IZ F

I

WWRIM.-

FLANKING SEQUENCET

\IBA

I

1S'l' I

ok

8MA-Rt

KOS NS-

S A,71 +-. X.

ok

1)14 kl-

_

-

..F

{

-.

_ _

-

4.X

-

-

.7

-

-

22

-

14 ki)-

.

..r

.111i

410 4.8

-

3.7

-

2.3

FIG. 2. (A) Southern blots of DNA from VP16 deletion mutants. Autoradiographic exposures of replicate Southern blots of mutant (8MA) and wild-type (KOS) DNA separated in 0.8% agarose gels are shown. Hybridization probes are underscored in the diagrams shown at left: coding sequence probe, 990-bp ApaI-SacIl fragment of pVP16-KOS; flanking sequence probe, 1.4-kb XbaI-PstI insert of pVP16RVASty. Restriction enzymes used to digest the blotted viral DNA samples are indicated above each set of gel lanes. The positions of DNA size markers on each gel appear to the right of the KOS samples. (B) Southern blots of VP16 revertant. Autoradiographic exposures of replicate Southern blots of 8MA revertant (8MA-R) DNA are shown; conditions are as described for panel A.

coding sequences (Fig. 2A, upper panel). This probe hybridized with single restriction fragments in the KOS DNA samples (BamHI, -9 kb; EcoRI, -13.4 kb) but failed to hybridize with DNA from the VP16 deletion isolate (8MA), as expected. A second blot was probed with the insert of pVP16RVASty that contains only HSV-1 sequences flanking the deletion/substitution mutation (Fig. 2A, lower panel). This probe hybridized with fragments in both the KOS and 8MA DNA samples, also as expected. The hybridizing BamHI fragment of 8MA was predicted to be 11.8 kb, because the ICP6-,-galactosidase insert of 8MA is 2.8 kb larger than the VP16 sequences of KOS that were replaced. Two unique-sized fragments were observed to hybridize in EcoRI-digested 8MA DNA (-8.1 and -5.1 kb), because the ICP6-3-galactosidase insert also contains two additional EcoRI sites. A third replicate blot was hybridized with a 3-kb EcoRI fragment spanning the P-galactosidase-coding sequences in pD6P. This probe hybridized with a single band of the expected size in both 8MA DNA samples but failed to

hybridize with KOS DNA (data not shown). The lengths of hybridizing DNA fragments observed for each of the triplicate blots corresponded very well with the predicted fragment sizes. We concluded, therefore, that the desired VP16 deletion/substitution mutant had been generated, replacing VP16-coding sequences of HSV-1 with the ICP6-p-galactosidase gene cassette. The mutant isolate designated 8MA was selected for further characterization. Reversion of the 8MA mutation with a wild-type allele of VP16. The transfection process used to generate the recombinant HSV-1 VP16 deletion mutant is inherently mutagenic. A genetic revertant of 8MA, restored for VP16-coding sequences, was constructed as a control for the possible introduction of adventitious mutations during the construction and isolation of 8MA. This revertant would be expected to exhibit wild-type growth properties, as long as no other discernable mutations were introduced during recombinational mutagenesis. The revertant was constructed by simply reversing the mutagenesis procedure. That is, purified 8MA

VOL. 66, 1992

virion DNA was cotransfected into 16-8 cells with a wildtype VP16 DNA fragment from pVP16-KOS, and virus progeny were screened for isolates that no longer stained positively for P-galactosidase. Southern blot hybridizations for three isolates were identical (data not shown); the results for one isolate, designated 8MA-R, are shown in Fig. 2B. As expected, the revertant exhibited the same hybridization profile as wild-type HSV-1 KOS DNA (Fig. 2A and B). Replication defect of VP16-deleted HSV-1: efficiency of plaquing. VP16 deletion mutants of HSV-1 can be propagated in 16-8 cells, as demonstrated by the isolation of 8MA. To determine whether the 8MA isolate could also replicate in cells that do not express VP16, the plaquing efficiencies of wild-type (KOS), VP16-deleted (8MA), and VP16-reverted (8MA-R) viruses were measured simultaneously on monolayers of 16-8 cells and Vero-A1-3 (A1-3) cells. A1-3 is a G418-resistant Vero cell-derived cell line that was subjected to the same procedures used to generate the 16-8 cell line, except that the VP16 expression vector was never introduced. In the results from one typical experiment, the ratio of plaques formed on monolayers of A1-3 versus 16-8 cells was 0.9 for KOS and 0.8 for 8MA-R. This indicated that the efficiency of plaquing for both KOS and 8MA-R was very similar in the two cell lines. An 8MA virus stock made in 16-8 cells produced a titer of 5 x 10' PFU/ml when measured on monolayers of 16-8 cells. This mutant virus stock was unable to form plaques on A1-3 cell monolayers, however, when as many as 105 PFU (titrated on 16-8 cells) were used as an inoculum. This corresponded to an MOI of approximately 0.1 PFU per cell. A 10-fold increase in MOI to 1.0 PFU per cell resulted in nearly confluent lysis of the A1-3 monolayer. Thus, the VP16 deletion mutant retained severe cytotoxicity for cells that do not express VP16, even though the efficiency of plaque formation was reduced by >105-fold in Al-3 cells. Identical results were also observed by using Vero cells infected with-8MA and several other VP16 deletion mutant isolates (data not shown). Replication defect of VP16-deleted HSV-1: efficiency of replication;. Th'e results of plaque assays described above indicate that the HSV-1 VP16 deletion/substitution mutant is unable to spread from cell to cell in A1-3 monolayers and that the revertant virus formed plaques like wild-type HSV-1. Intracellular replication of the VP16 deletion mutant was examined by establishing single-step replication curves for KOS, 8MA, and 8MA-R in both 16-8 and A1-3 cells, using an MOI of 5 PFU per cell. Replicate plates from each infected culture were harvested at various times postinfection and assayed for infectious virus by plaque titration on 16-8 cells. All three viruses replicated efficiently in 16-8 cells, and the time course of replication was the same for each virus (Fig. 3, upper graph). Only KOS and 8MA-R replicated in A1-3 cells, however; replication of the 8MA mutant was not detectable (Fig. 3, lower graph). Taken together, data from the plaque assays and singlestep replication studies described above indicate that the VP16 deletion mutant of HSV-1 is unable to replicate in cells that do not express VP16. This defect was complemented efficiently, however, in cells that are transformed with a VP16 expression vector (16-8 cells). Moreover, since the revertant virus displayed wild-type replication properties, neither it nor 8MA harbors additional mutations that effect viral replication in tissue culture cells. VP16 protein in 8MA virions. The possibility was considered that VP16 expressed by 16-8 cells might be recruited to serve as a structural component of 8MA virions during the

VP16 DELETION MUTANT OF HSV-1

263

10 9

16-8 CELLS

107.

10 .

S

10

vl4o 10 a-

g

A1-3 CELLS

,8.

107 lo6gI 16. lo

-

- -

-

.

S

0

10

20

30

40

HOURS POSTINFECTION

FIG. 3. Single-step growth

curves

in 16-8 and A1-3 cells. Repli-

cate six-well cluster dishes of 16-8 (upper graph) and A1-3 (lower

graph) cells were infected with KOS, 8MA, or 8MA-R (MOI, 5 PFU per cell). Individual culture wells were harvested at time points from 2 to 36 h postinfection (x axis) by scraping infected cells into the culture medium and subjecting this suspension to three freeze-thaw cycles (on dry ice and at room temperature). The total yield of PFU at each time point was determined by plaque titration on 16-8 cells (y axis). Solid line, KOS; dashed line, 8MA; dotted line, 8MA-R.

propagation of the mutant. Western immunoblots of purified virions were carried out to determine the level of VP16 in 8MA virions relative to that in KOS and 8MA-R virions. Equivalent PFU amounts of each infectious virus were subjected to SDS-PAGE, transferred to nitrocellulose filters, and probed with a mixture of anti-VP16 amino and carboxyl terminal peptide antisera (Fig. 4; see Materials and Methods). The relative amount of immunoreactive VP16 per PFU was determined by measuring the absorbance of the stained VP16 bands with a laser densitometer. In the blot shown (Fig. 4), 8MA virions contained from 1.6 to 1.9 times more VP16 per PFU than did KOS virions and 2.3 to 4.9 times more than did 8MA-R virions. Similar data were observed for Western blots of crude virus stocks from one additional VP16 deletion isolate and its revertant (data not shown). We concluded that, as a result of propagation in 16-8 -cells, cell-derived VP16 was recruited by the 8MA mutant to serve as a structural component of deletion mutant virions. Plasmid transactivation assays were performed to confirm

264

J. VIROL. WEINHEIMER ET AL.J.Vo. VP16A MUTANTF 8MA 3.2 x

3.2 x

to 4 10

10

106

REVERTANT' 8NIA-R

WILD-TYPE KOS

3.2 x

3.2 x

1 05

105 1i6 i05 10, 106

PF' -

200

I110 kI)

-84

-47

FIG. 4. VP16 immunoblots of HSV-1 virions. 8MA, KOS, and 8MA-R virion preparations (indicated at top) were diluted to obtain aliquots containing a range of PFU totals (indicated above each gel lane). Virion proteins were separated on 4 to 20% SDS-PAGE gels, transferred to nitrocellulose, and probed with a mixture of anti-VP16 peptide antisera (SW1 plus SW7; see Materials and Methods). The positions of protein molecular mass markers are shown at right. The position of immunoreactive VP16 is indicated with an arrow.

that the vinion-associated VP16 of 8MA functions normally for the induction of IE gene transcription. Cells transfected with various promoter plasmids were superinfected with 8MA, 8MA-R and KOS in the presence of 100 p.g of cycloheximide per ml. IE promoters were specifically transinduced by all three viruses; DE, late, and non-HSV promoter plasmids, however, were not trans-induced (4b). In a second set of experiments using cells that were simply infected without prior transfection, Western immunoblots carried out over a time course of infection confirmed that IE proteins are produced at very similar levels in Al-3 and 16-8 cells infected with KOS, 8MA, or 8MA-R (4a). We concluded from these data that the VP16 protein associated with 8MA virions carries out the normal regulatory function expected of VP16. Viral DNA replication. At what point during infection does the absence of VP16 protein synthesis restrict lytic replication? In a typical HSV-1 infection, the synthesis of functional IE proteins leads to DE gene expression and, in turn, to viral DNA replication. We examined the efficiency of viral DNA replication by DNA hybridization over a time course of infection with KOS, 8MA, and 8MA-R in both Al-3 and 16-8 cells. For all three viruses, no difference was observed between the DNA replication profiles of Al-3 and 16-8 cells (Fig. 5). Moreover, the VP16 deletion mutant, 8MA, induced normal levels of viral DNA synthesis through 9 h postinfection in both cell lines. The accumulation of mutant viral DNA was slightly reduced relative to that of wild-type and revertant viruses at 12 h postinfection in both 16-8 and Al-3 cells. This result was reproducible in 16-8, Al-3, and Vero cells (additional data not shown). The observed difference does not likely result from the absence of VP16 protein synthesis per se, as it was seen in both VP16-transformed (16-8) and normal (Al-3 and Vero) cells (Fig. 5; additional data not shown). The abnormality was corrected, however, in the VP16 revertant, 8MA-R, so it is specific to the deletion/substitution allele of 8MA. Nevertheless, the time course of 8MA DNA replication was essentially identical in 16-8 and Al-3 cells. This fact indicates that the restriction point for the replication of the HSV-1 VP16 deletion mutant in Al-3 cells occurs at a step that is independent of the requireme'nts for efficient HSV-1 DNA replication. HSV-1 DNA encapsidation. Efficient HSV-1 DNA replica-

HOURS POSTINFECTION FIG. 5. HSV-1 DNA synthesis. Replicate six-well culture dishes of 16-8 (upper graph) and Al-3 (lower graph) cells were infected with KOS, 8MA, or 8MA-R (MOI, 10 PFU per cell). Individual culture wells were harvested between 2 and 24 h postinfection (x axis), and DNA was extracted and normalized for the recovery of a tritiumlabeled Vero cell DNA standard added at the time of cell lysis. The normalized counts per minute of radiolabeled hybridized HSV-1 DNA (see Materials and Methods) is plotted on the y axis. Solid line, KOS; dashed line, 8MA; dotted line, 8MA-R.

tion normally leads to induced levels of late viral protein synthesis, viral DNA encapsidation, and then virion assembly. During this process, concatemeric HSV-1 chromosomal DNA is cleaved into chromosome-length fragments (9, 18, 22) (Fig. 6). Consequently, encapsidated viral DNA contains free chromosomal termini whereas nonpackaged concatemeric DNA does not. Thus, the ratio of cleaved to concatemeric terminal DNA in total cell lysates can be used as an indirect measure of the level of encapsidation (3). This ratio can be monitored readily by restriction enzyme digestion and Southern blotting, as in the experiment described below. Infected cell lysates were prepared from 16-8 and Al-3 cells infected with KOS, 8MA, or 8MA-R. Total DNA was purified and digested with BamHI restriction endonuclease. BamHl cuts in the repeated sequences that occur at either end of the HSV-1 chromosome and at the junction of the long and short genome segments (Fig. 6). Only encapsidated DNA with free chromosomal termini will give rise to the BamHl S and P fragments of HSV-1 DNA. Concatemeric HSV-1 DNA, on the other hand, gives rise only to the

VOL. 66, 1992

VP16 DELETION MUTANT OF HSV-1

t /~~~~~~~~~

CHROMOSOMAL

UN1rr

~II1

BAM HI

t

-m//

BAM HI BAM HI

H

BAM HI

H

SH

s

265

p FIG. 6. HSV-1 terminal sequences. Concatemeric HSV-1 DNA from which single chromosomes (between arrows) are cleaved during encapsidation is depicted. Open boxes indicate the terminal (TRL and TRs) and inverted internal (IRL and IRS) repeats of HSV-1 DNA. The locations of relevant BamHI restriction enzyme sites in the repeat units, with the identities of resulting BamHI restriction fragments (S, P, and SP), are indicated.

BamHI SP fragment. BamHI digestion products were separated by agarose gel electrophoresis, Southern blotted, and probed with radiolabeled BamHI SP DNA to detect all forms of the terminal sequences. The blots were probed simultaneously with radiolabeled BamHI B fragment of HSV-1 DNA, which does not hybridize with terminal DNA sequences, so that hybridization signals for the BamHI S, P, and SP fragments in each sample could be normalized for chromosome number (see Materials and Methods). The ratio of hybridization signals for the S plus P fragments to those for the SP fragments was calculated in order to obtain an indirect measure of HSV-1 DNA encapsidation. In two trials the extent of viral DNA encapsidation appeared to be similar (74 to 135% of KOS levels) in 16-8 cells infected with KOS, 8MA, or 8MA-R and in A1-3 cells infected with either KOS or 8MA-R (Table 1). Lower levels of viral DNA encapsidation (30 to 54% of KOS levels) in A1-3 cells infected with 8MA were detected (Table 1). One-half of each infected cell lysate was treated with DNase I prior to DNA extraction and was then processed alongside the untreated samples as a control to validate measurements of encapsidation. DNase-treated samples should consist only of encapsidated HSV-1 DNA that was protected from nuclease digestion. This would be expected to result in the same ratio of S plus P fragments to SP fragments for all samples, and this was the case to within ±+17% (data not shown). We concluded from our analysis that viral DNA encapsidation is probably reduced by at least 50 to 75% as a result of the VP16 deletion. Though significant, this percent reduction in DNA encapsidation did not appear to account for the lethal replication defect caused by the VP16 deletion mutation. TEM. The accumulation of virus particles during wildtype and mutant HSV-1 infection was examined directly by TEM to determine whether the replication defect of 8MA might be associated with the process of virion assembly. Cultures of 16-8 and A1-3 cells were fixed and prepared for TEM after 16 h of infection with KOS, 8MA, or 8MA-R. The abundance of viral capsid forms (Fig. 7) in the nucleus and cytoplasm of each infected culture was tabulated (Table 2). Infections of 16-8 cells with KOS, 8MA, or 8MA-R and infections of A1-3 cells with either KOS or 8MA all gave very similar profiles under TEM. High numbers of virus particles appeared in both the nucleus and cytoplasm of infected cells, and the predominant capsid form in both populations had dense cores indicative of encapsidated viral DNA (Table 2 and Fig. 7). Cytoplasmic capsids in all of these samples were predominantly enveloped (Fig. 7). Enveloped, cytoplasmic particles often appeared in groups, often in association with membranous organelles, and were often

SP

localized at the cell surface or in the extracellular space (Fig. 7). These productive infections were characterized further by ultrastructural pathology characteristic of productive HSV-1 infections (nuclear membrane distortion and reduplication, chromatin margination, nucleolar segregation, etc.) (reviewed in reference 8 and data not shown). Slightly reduced total numbers of virus capsids (-50%) were observed for A1-3 cells infected with 8MA. In contrast to the previous cases, 8MA capsids in A1-3 cells were predominantly empty or partially cored (Table 2 and Fig. 7), although they were similarly distributed in the nucleus and cytoplasm and were predominantly enveloped in the cytoplasm. The dense-cored particles that did appear in the cytoplasm were individual, and they did not accumulate at the cell surface or in the extracellular space. An additional population of heterogeneous aberrant capsid forms appeared in A1-3 cells infected with 8MA much more frequently than in any of the productively infected cell samples (approximately 10-fold; data not shown). Finally, the overt subcellular pathology characteristic of productive infection was not present in A1-3 cells infected with 8MA (data not shown). We concluded from this study that the elimination of VP16 protein synthesis during HSV-1 infection leads to a profound defect in the assembly of infectious virions. DISCUSSION A VP16 deletion mutant of HSV-1 was constructed and propagated in a cell line transformed with a VP16 expression vector. VP16 produced by this cell line was recruited into virions of the deletion mutant during propagation. This allowed IE gene expression to be induced normally during subsequent infection with the mutant virions, even in cells that do not express VP16. During infection of such a cell line (A1-3 cells), 8MA DNA replication and capsid formation were shown to occur at nearly normal levels. It was also shown that the level of 8MA DNA encapsidation was slightly TABLE 1. HSV-1 DNA encapsidation Cleavage ratio (cpm of S 16-8 cells

Virus

KOS 8MA 8MA-R "

+

P/cpm of SP)' A1-3 cells

Expt 1

Expt 2

Expt 1

0.34 (100) 0.25 (74) 0.37 (109)

0.26 (100) 0.27 (104) 0.35 (135)

0.14 (30) 0.34 (74)

0.46 (100)

Expt 2 0.63 (100) 0.34 (54) 0.51 (81)

Percentage relative to the value for KOS given in parentheses.

266

.,

J. VIROL.

WEINHEIMER ET AL.

A

0

.O. 1+

z?

.

#.

...

dC

..

I? Owl

t -1

f

, w

v

at

ll. e.

sr

(D .r

i. A

I

__

1-

.^

tF'

0

-1.-h -. -

.4'

.r

G6iitr 'O

I

.%,41.

xS

*1 jl:. I

N

I.

4'...

3.

'K-

wt t,~~~~ 4~ ~ ~

~

~

~~~~~~~V .-

4:

4 k

-

.n

C.

0

FIG. 7. Transmission electron micrographs of HSV-1 particles forms. (A) Infection of A1-3 cells with 8MA (magnification, x30,240). There are several empty or partial nucleocapsids (enclosed in circles) in the nucleus. A single empty or partial enveloped capsid is present in the cytoplasm (arrow). (B) Infection of 16-8 cells with 8MA (magnification, x30,240). Numerous dense enveloped capsids (arrows) are adjacent to the nuclear envelope and in the extracellular space. A dense capsid is present in the nucleus (enclosed in circle). (C) Infection of A1-3 cells with 8MA-R (magnification, x 30,240). Dense enveloped capsids (enclosed in circles) are present in the cytoplasm and extracellular space. A dense particle is budding from the reduplicated nuclear membrane (arrow).

VOL. 66, 1992

VP16 DELETION MUTANT OF HSV-1

267

TABLE 2. HSV-1 particle forms observed by TEM Virus/cell type

Nucleus

Abundance of capsid forms in": Cytoplasm and/or extracellular space

Total capsids

Empty/partial

Dense

Empty/partial

Dense

KOS/16-8 8MA/16-8 8MA-R/16-8

6.0 (100) 5.6 (93) 5.3 (88)

13.0 (100) 10.0 (77) 9.7 (75)

5.8 (100) 5.3 (91) 5.0 (86)

16.6 (100) 17.3 (104) 15.0 (90)

41.4 (100) 38.2 (92) 35.0 (85)

KOS/A1-3 8MA/A1-3 8MA-R/A1-3

5.5 (100) 8.0 (145) 5.0 (91)

11.0 (100) 1.8 (16) 8.5 (100)

5.0 (100) 7.5 (150) 4.0 (80)

15.5 (100) 1.0 (6) 15.5 (100)

37.0 (100) 18.3 (49) 33.0 (89)

a Data expressed as the mean number of capsids per cell, with the percentage relative to the value for KOS given in parentheses.

reduced in A1-3 cells relative to wild-type and revertant controls. Consistent with this observation, a reduction in the level of dense-cored capsids produced by 8MA was also observed by TEM. The process of 8MA virion assembly appeared to be profoundly disturbed in A1-3 cells, and the production of infectious 8MA virions could not be detected by either plaque assay or single-step replication assay. Together, these data indicate that the VP16 deletion mutation in 8MA produced a lethal phenotype that correlates with a defect in virion assembly. Specificity of the phenotype observed for 8MA was addressed in two ways. First, the defect was complemented efficiently in 16-8 cells that express VP16. VP16 is the only known protein product of the HSV-1 sequences present in 16-8 cells (5, 7, 16, 26, 29), and 16-8 cells were also shown to complement the HSV-2 mutant ts2203, consistent with a previous report that HSV-1 DNA fragments encoding VP16 rescue the ts2203 mutation (1). The second test of specificity used a site-specific revertant of the deletion mutant that was restored for VP16-coding sequences. The revertant exhibited fully wild-type lytic replication properties, indicating that the replication defect of 8MA mapped entirely within the DNA fragment used to rescue the mutation, from 455 bp upstream of the VP16 ORF initiation codon to 944 bp downstream of the VP16 termination codon. All of the phenotypic properties exhibited by 8MA were shown to be specific for VP16 functions, with the likely exception of a slight reduction in the level of DNA replication observed at 12 h postinfection for both A1-3 and 16-8 cells. This reduction could be due to the loss of VP16 function, but only if the 16-8 cell-derived VP16 is altered or modified in a way that prevents full complementation of the DNA replication pattern. Alternatively, a mutation in sequences immediately flanking the VP16 ORF might indicate a secondary gene function required for maximal DNA replication. The use of an expressing cell line to propagate mutants of a viral structural protein leads to the production of virions bearing a mutated gene on the encapsidated chromosome and wild-type structural protein in the surrounding virion. The observation that VP16 protein expressed by 16-8 cells can be recruited into 8MA virions suggests that the cellderived protein functions normally in HSV-1 virion assembly. This conclusion is not definitive, however, pending viral particle counts to determine the relative number of VP16 molecules per virion of the mutant and wild-type strains propagated in 16-8 cells. Slightly higher levels of VP16 per PFU of 8MA than per PFU of KOS and 8MA-R were detected, and the potential effects of increased amounts of VP16 per PFU are unknown. Because of the ample supply of VP16 per PFU of 8MA virions, however, 8MA induced IE

gene expression efficiently in cells that do not express VP16. This latter observation indicates that the cell-derived, virionassociated VP16 also functions normally in IE gene transinduction. The introduction of this functional VP16 during the infection of A1-3 cells creates the potential for further phenotypic leakage, and this must be considered when analyzing the effects of the mutation. The infection of cells not expressing VP16 with the VP16 deletion mutant allows a new opportunity to examine the fate of VP16 that is introduced with infecting virions. In this report, we have shown that the VP16 is functionally intact but is not recycled efficiently for another round of virion assembly, since less than 1% of 8MA input infectivity can be detected at late times of infection in A1-3 cells. The input VP16 is stable during the infection of A1-3 cells, since Western immunoblot analyses have revealed no apparent change in the amount or electrophoretic migration of VP16 during the first 24 h of infection (4a). The efficient assembly of virions must require much larger amounts of available, and perhaps freshly synthesized, VP16. It is possible that the virion-associated VP16 is either altered or sequestered in some way during infection so that it is unable to serve again as a virion structural component. Very low levels of dense-cored capsids in the cytoplasm of A1-3 cells infected with 8MA were observed, but they apparently are not infectious. These cytoplasmic particles might be attributable to residual VP16 function provided by the input VP16 from 8MA virions, as this protein probably retains some functional activity. Alternatively, these particles might arise from the same processes that also give rise to larger numbers of abherent capsid forms. In either case, these particles are most likely defective, since their appearance does not correlate with the detection of infectious 8MA progeny. 8MA DNA replication and capsid formation were shown to occur at nearly normal levels in A1-3 cells. Slightly reduced levels of 8MA DNA encapsidation were measured by hybridization in A1-3 cells, and this level was higher than but still consistent with the reduced level of dense-cored capsids observed by TEM. It is not clear why viral DNA encapsidation or the accumulation of intranuclear densecored capsids would be negatively affected by the loss of VP16 function. Although this finding might appear to implicate VP16 directly in DNA encapsidation, an indirect effect of the VP16 mutation might also produce this phenotype and should not be ruled out (see below). The VP16 deletion mutant expresses normal levels of IE proteins and also produces normal levels of replicated viral DNA through 9 h postinfection. It would be expected from these observations that DE protein synthesis occurs nor-

268

WEINHEIMER ET AL.

mally, since DE proteins are induced by IE proteins and are required for HSV-1 DNA replication. HSV DNA replication normally provides, in turn, the inductive stimulus for high levels of late protein synthesis. By examining protein synthesis patterns, we have observed that DE and late proteins are synthesized at the appropriate time intervals during the infection of A1-3 and Vero cells with the 8MA virus (4b). This suggests that the temporal regulation of protein synthesis is intact in the absence of VP16 protein synthesis during infection. Quantitatively, however, the synthesis of all HSV-1 proteins at later times is significantly reduced in cells that do not express VP16. This quantitative defect is complemented efficiently in 16-8 cells (4b). Manifestation of a mutant phenotype late in infection might be expected from the loss of newly synthesized VP16, since VP16 is a late gene product. It is surprising, however, that the synthesis of other late proteins is reduced as well, since no regulatory function other than IE trans-induction has been ascribed to VP16. It is also not clear what effect low levels of viral protein synthesis at later times of infection might have on concurrent processes like DNA encapsidation or virion assembly. VP16 is the first HSV-1 tegument protein to be assigned an essential role in HSV-1 lytic replication. In the case of two other known tegument proteins, UL47 (27) and US9 (10), the ORFs of the respective proteins are not required for HSV-1 replication (32, 45). The mechanistic details of HSV-1 virion assembly are poorly understood, and a functional role for VP16 in virion assembly has not been defined clearly, nor has this study completely elucidated the role of VP16. It is possible that VP16 plays an important structural role in the virion, perhaps as a tether between tegument proteins themselves or between tegument proteins and the virion capsid or envelope proteins, or both. In this study, the overwhelming shift from the production of dense-cored capsids to the production of aberrant, empty-, and partially cored capsids in A1-3 cells infected with 8MA might result from the loss of this function. The failure to produce infectious progeny could, in turn, result directly from this assembly defect. An essential role of VP16 for HSV-1 virion assembly is a second key function of VP16 during HSV-1 infection. VP16 trans-induction of IE gene transcription during HSV-1 infection and the role of this activity in viral pathogenesis have been documented previously (2, 39). It will be of particular interest to define more precisely the function of VP16 during HSV-1 virion morphogenesis. Moreover, it will be of interest to

compare

the structural features of VP16 required for

transactivation (1, 15, 41, 43) with those required for virion morphogenesis. Critical sites on VP16 that affect both of these functions have been defined by mutagenesis, whereas other sites specifically affect only one of these functions (1, 42a). The availability of site-specific mutants and their future introduction into the HSV-1 chromosome might help to characterize more precisely the functions, and functional domains, of VP16 in both gene trans-induction and virion assembly. ACKNOWLEDGMENTS

We are grateful to V. Preston for providing ts2203 and to S. Triezenberg, S. McKnight, S. Weller, R. Sandri-Goldin, and B. Roizman for plasmids used in this work. Nick Bahia provided excellent technical assistance for TEM. We thank A. K. Field for support, F. Tufaro for advice

discussions, and A. comments

on

on

electron microscopy and for helpful

K. Field, R. Hamatake, and C. Dilanni for

the manuscript.

J. VIROL.

REFERENCES 1. Ace, C. I., M. A. Dalrymple, F. H. Ramsay, V. G. Preston, and C. M. Preston. 1988. Mutational analysis of the herpes simplex virus type 1 trans-inducing factor, Vmw65. J. Gen. Virol. 69:2595-2605. 2. Ace, C. I., T. A. McKee, J. M. Ryan, J. M. Cameron, and C. M. Preston. 1989. Construction and characterization of a herpes simplex virus type 1 mutant unable to transinduce immediateearly gene expression. J. Virol. 63:2260-2269. 3. Al-Kobaisi, M. F., F. J. Rixon, I. McDougall, and V. G. Preston. 1991. The herpes simplex virus UL33 gene product is required for the assembly of full capsids. Virology 180:380-388. 4. Batterson, W., and B. Roizman. 1983. Characterization of the herpes simplex virion-associated factor responsible for the induction of a genes. J. Virol. 46:371-377. 4a.Boyd, B., D. O'Boyle, and S. Weinheimer. Unpublished data. 4b.Boyd, B., and S. Weinheimer. Unpublished data. 5. Campbell, M. E. M., J. W. Palfreyman, and C. M. Preston. 1984. Identification of herpes simplex virus DNA sequences which encode a trans-acting polypeptide responsible for stimulation of immediate-early transcription. J. Mol. Biol. 180:1-19. 6. Corsaro, C. M., and M. L. Pearson. 1981. Enhancing the efficiency of DNA-mediated gene transfer in mammalian cells. Somatic Cell Genet. 7:603-616. 7. Dalrymple, M. A., D. J. McGeoch, A. J. Davison, and C. M. Preston. 1985. DNA sequence of the herpes simplex virus type 1 gene whose product is responsible for transcriptional activation of immediate early promoters. Nucleic Acids Res. 13:78657879. 8. Dargan, D. J. 1986. The structure and assembly of herpesviruses, p. 359-437. In J. Harris and R. Horne (ed.), Electron microscopy of proteins, vol. 5. Academic Press, Inc. (London), Ltd., London. 9. Deiss, L. P., and N. Frenkel. 1986. Herpes simplex virus amplicon: cleavage of concatemeric DNA is linked to packaging and involves amplification of the terminally reiterated a sequence. J. Virol. 57:933-941. 10. Frame, M. C., D. J. McGeoch, F. J. Rixon, A. C. Orr, and H. S. Marsden. 1986. The 10K virion phosphoprotein encoded by gene US9 from herpes simplex virus type 1. Virology 150:321332. 11. Gerster, T., and R. G. Roeder. 1988. A herpesvirus transactivator protein interacts with transcription factor OTF-1 and other cellular proteins. Proc. Natl. Acad. Sci. USA 85:6347-6351. 12. Goldin, A. L., R. M. Sandri-Goldin, M. Levine, and J. C. Glorioso. 1981. Cloning of herpes simplex virus type 1 sequences representing the whole genome. J. Virol. 38:50-58. 13. Goldstein, D. J., and S. K. Weller. 1988. An ICP6::lacZ insertional mutagen is used to demonstrate that the UL52 gene of herpes simplex virus type 1 is required for virus growth and DNA synthesis. J. Virol. 62:2970-2977. 14. Graham, F. L., and A. J. van der Eb. 1973. A new technique for the assay of infectivity of human adenovirus 5 DNA. Virology 52:456-467. 15. Greaves, R., and P. O'Hare. 1989. Separation of requirements for protein-DNA complex assembly from those for functional activity in the herpes simplex virus regulatory protein Vmw65. J. Virol. 63:1641-1650. 16. Hall, L. M., K. G. Draper, R. J. Frink, R. H. Costa, and E. K. Wagner. 1982. Herpes simplex virus mRNA species mapping in EcoRl fragment I. J. Virol. 43:594-607. 17. Honess, R. W., and B. Roizman. 1974. Regulation of herpesvirus macromolecular synthesis. I. Cascade regulation of the synthesis of three groups of viral proteins. J. Virol. 14:8-19. 18. Jacob, R. J., L. S. Morse, and B. Roizman. 1979. Anatomy of herpes simplex virus DNA. XII. Accumulation of head to tail concatemers in nuclei of infected cells and their role in the generation of the four isomeric arrangements of viral DNA. J. Virol. 29:448-457. 19. Kristie, T. M., J. H. LeBowitz, and P. A. Sharp. 1989. The octamer-binding proteins form multi-protein-DNA complexes with the HSV aTIF regulatory protein. EMBO J. 8:4229-4238. 20. Kristie, T. M., and B. Roizman. 1988. Differentiation and DNA

VOL. 66, 1992

21. 22.

23.

24.

25. 26.

27.

28.

29.

30.

31.

32.

contact points of host proteins binding to the cis site for virion-mediated induction of a genes of herpes simplex virus 1. J. Virol. 62:1145-1157. Kristie, T. M., and P. A. Sharp. 1990. Interactions of the Oct-1 POU subdomains with specific DNA sequences and with the HSV a-trans-activator protein. Genes Dev. 4:2383-23%. Ladin, B. F., S. Ihara, H. Hampl, and T. Ben-Porat. 1982. Pathway of assembly of herpesvirus capsids: an analysis using DNA' temperature sensitive mutants of pseudorabies virus. Virology 116:544-561. Lemaster, S., and B. Roizman. 1979. Herpes simplex virus phosphoproteins. II. Characterization of the virion protein kinase and of the polypeptides phosphorylated in the virion. J. Virol. 35:798-811. Mackem, S., and B. Roizman. 1982. Structural features of the herpes simplex virus ax gene 4, 0, and 27 promoter-regulatory sequences which confer a regulation on chimeric thymidine kinase genes. J. Virol. 44:939-949. Marsden, H. S., N. D. Stow, V. G. Preston, M. C. Timbury, and N. M. Wilkie. 1978. Physical mapping of herpes simplex virusinduced polypeptides. J. Virol. 28:624-42. McGeoch, D. J., M. A. Dalrymple, A. J. Davison, A. Dolan, M. C. Frame, D. McNab, L. J. Perry, J. E. Scott, and P. Taylor. 1988. The complete DNA sequence of the long unique region in the genome of herpes simplex virus type 1. J. Gen. Virol. 69:1531-1574. McLean, G., F. Rixon, N. Langeland, L. Haarr, and H. Marsden. 1990. Identification and characterization of the virion protein products of herpes simplex virus type 1 gene UL47. J. Gen. Virol. 71:2953-2960. O'Hare, P., and C. R. Goding. 1988. Herpes simplex virus regulatory elements and the immunoglobulin octamer domain bind a common factor and are both targets for virion transactivation. Cell 52:435-445. Pellett, P. E., J. L. C. McKnight, F. J. Jenkins, and B. Roizman. 1985. Nucleotide sequence and predicted amino acid sequence of a protein encoded in a small herpes simplex virus DNA fragment capable of trans-inducing a genes. Proc. Natl. Acad. Sci. USA 82:5870-5874. Post, L. E., A. J. Conley, E. S. Mocarski, and B. Roizman. 1980. Cloning of reiterated and nonreiterated herpes simplex virus 1 sequences as Bam HI fragments. Proc. Natl. Acad. Sci. USA 77:4201-4205. Post, L. E., S. Mackem, and B. Roizman. 1981. Regulation of a genes of herpes simplex virus: expression of chimeric genes produced by fusion of thymidine kinase with a gene promoters. Cell 24:555-565. Post, L. E., and B. Roizman. 1981. A generalized technique for deletion of specific genes in large genomes: a gene 22 of herpes

VP16 DELETION MUTANT OF HSV-1

269

simplex virus is not essential for growth. Cell 25:227-232. 33. Preston, C. M., M. C. Frame, and M. E. M. Campbell. A complex formed between cell components and an HSV structural polypeptide binds to a viral immediate early gene regulatory DNA sequence. Cell 52:425-434. 34. Rigby, P. W. J., M. Dieckmann, C. Rhodes, and P. Berg. 1977. Labelling of DNA to high specific activity by nick translation. J. Mol. Biol. 113:237-258. 35. Roizman, B., and F. J. Jenkins. 1985. Genetic engineering of novel genomes of large DNA viruses. Science 229:1208-1214. 36. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 37. Southern, E. M. 1975. Detection of specific sequences among DNA fragments separated by gel electrophoresis. J. Mol. Biol. 98:503-517. 38. Southern, P. J., and P. Berg. 1982. Transformation of mammalian cells to antibiotic resistance with a bacterial gene under control of the SV40 early region promoter. J. Mol. Appl. Genet. 1:327-341. 39. Steiner, I., J. G. Spivak, S. L. Deshmane, C. I. Ace, C. M. Preston, and N. W. Fraser. 1990. A herpes simplex virus type 1 mutant containing a nontransinducing Vmw65 protein establishes latent infection in vivo in the absence of viral replication and reactivates efficiently from explanted trigeminal ganglia. J. Virol. 64:1630-1638. 40. Stern, S., M. Tanaka, and W. Herr. 1989. The Oct-1 homoeodomain directs formation of a multiprotein-DNA complex with HSV transactivator VP16. Nature (London) 341:624-630. 41. Triezenberg, S. J., R. C. Kingsbury, and S. L. McKnight. 1988. Functional dissection of VP16, the trans-activator of herpes simplex virus immediate early gene expression. Genes Dev. 2:718-729. 42. Wagner, E. K. 1985. Individual HSV transcripts: characterisation of specific genes, p. 45-104. In B. Roizman (ed.), The herpesviruses, vol. 3. Plenum Publishing Corp., New York. 42a.Weinheimer, S., D. O'Boyle, and B. Boyd. Unpublished data. 43. Werstuck, G., and J. P. Capone. 1989. Mutational analysis of the herpes simplex virus trans-inducing factor Vmw65. Gene 75: 213-224. 44. Xiao, P., and J. P. Capone. 1990. A cellular factor binds to the herpes simplex virus type 1 transactivator Vmw65 and is required for Vmw65-dependent protein-DNA complex assembly with Oct-1. Mol. Cell. Biol. 10:4974-4977. 45. Zhang, Y., D. A. Sirko, and J. L. C. McKnight. 1991. Role of herpes simplex virus type 1 UL46 and UL47 in aTIF-mediated transcriptional induction: characterization of three viral deletion mutants. J. Virol. 65:829-841.