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Simultaneous Deletion of the 9GL and UK Genes from the African Swine Fever Virus Georgia 2007 Isolate Offers Increased Safety and Protection against Homologous Challenge

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Simultaneous Deletion of the

9GL

and

UK

Genes from the African Swine Fever

Virus Georgia 2007 Isolate Offers

Increased Safety and Protection against

Homologous Challenge

Vivian O’Donnell,a,bGuillermo R. Risatti,bLauren G. Holinka,aPeter W. Krug,a Jolene Carlson,a,cLauro Velazquez-Salinas,aPaul A. Azzinaro,aDouglas P. Gladue,a Manuel V. Borcaa

Agricultural Research Service, U.S. Department of Agriculture, Plum Island Animal Disease Center, Greenport,

New York, USAa; Department of Pathobiology and Veterinary Science, CANR, University of Connecticut, Storrs,

Connecticut, USAb; Biosecurity Research Institute and Department of Diagnostic Medicine and Pathobiology,

College of Veterinary Medicine, Kansas State University, Manhattan, Kansas, USAc

ABSTRACT African swine fever virus (ASFV) is the etiological agent of a contagious and often lethal viral disease of domestic pigs that has significant economic conse-quences for the swine industry. The control of African swine fever (ASF) has been hampered by the unavailability of vaccines. Successful experimental vaccines have been derived from naturally occurring, cell culture-adapted, or genetically modified live attenuated ASFV. Recombinant viruses harboring engineered deletions of spe-cific virulence-associated genes induce solid protection against challenge with pa-rental viruses. Deletion of the 9GL(B119L) gene in the highly virulent ASFV isolates Malawi Lil-20/1 (Mal) and Pretoriuskop/96/4 (Δ9GL viruses) resulted in complete pro-tection when challenged with parental isolates. When similar deletions were created within the ASFV Georgia 2007 (ASFV-G) genome, attenuation was achieved but the protective and lethal doses were too similar. To enhance attenuation of ASFV-G, we deleted another gene,UK(DP96R), which was previously shown to be involved in at-tenuation of the ASFV E70 isolate. Here, we report the construction of a double-gene-deletion recombinant virus, ASFV-G-Δ9GL/ΔUK. When administered intramuscu-larly (i.m.) to swine, there was no induction of disease, even at high doses (106

HAD50). Importantly, animals infected with 104 50% hemadsorbing doses (HAD50) of

ASFV-G-Δ9GL/ΔUK were protected as early as 14 days postinoculation when chal-lenged with ASFV-G. The presence of protection correlates with the appearance of serum anti-ASFV antibodies, but not with virus-specific circulating ASFV-specific gamma interferon (IFN-␥)-producing cells. ASFV-G-Δ9GL/ΔUK is the first rationally de-signed experimental ASFV vaccine that protects against the highly virulent ASFV Georgia 2007 isolate as early as 2 weeks postvaccination.

IMPORTANCE Currently, there is no commercially available vaccine against African swine fever. Outbreaks of the disease are devastating to the swine industry and are caused by circulating strains of African swine fever virus. Here, we report a putative vaccine derived from a currently circulating strain but containing two deletions in two separate areas of the virus, allowing increased safety. Using this genetically modified virus, we were able to vaccinate swine and protect them from developing ASF. We were able to achieve protection from disease as early as 2 weeks after vac-cination, even when the pigs were exposed to a higher than normal concentration of ASFV.

KEYWORDS 9GL, ASFV, African swine fever virus,UK, vaccine

Received12 September 2016Accepted18 October 2016

Accepted manuscript posted online26 October 2016

CitationO'Donnell V, Risatti GR, Holinka LG, Krug PW, Carlson J, Velazquez-Salinas L, Azzinaro PA, Gladue DP, Borca MV. 2017. Simultaneous deletion of the9GLandUK

genes from the African swine fever virus Georgia 2007 isolate offers increased safety and protection against homologous challenge. J Virol 91:e01760-16. https://doi.org/10.1128/ JVI.01760-16.

EditorStanley Perlman, University of Iowa

Copyright© 2016 American Society for Microbiology. All Rights Reserved.

Address correspondence to Manuel V. Borca, manuel.borca@ars.usda.gov.

crossm

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A

frican swine fever (ASF) is a contagious and often fatal viral disease of swine. The causative agent, ASF virus (ASFV), is a large enveloped virus containing a double-stranded DNA (dsDNA) genome of approximately 190 kbp (1). ASFV shares aspects of genome structure and replication strategy with other large dsDNA viruses, including the familiesPoxviridae,Iridoviridae, andPhycodnaviridae(2).

Currently, the disease is endemic in more than 20 sub-Saharan African countries. In Europe, ASF is endemic on the island of Sardinia (Italy), and new outbreaks have been declared in the Caucasus region since 2007, affecting Georgia, Armenia, Azerbaijan, and Russia. Isolated outbreaks have been recently reported in Ukraine, Belarus, Lithuania, Latvia, and Poland, posing the risk of further dissemination into neighboring countries. The epidemic virus, ASFV Georgia 2007/1 (ASFV-G), is a highly virulent isolate belonging to the ASFV genotype II group (3).

Currently, there is no vaccine available for ASF, and disease outbreaks are controlled by animal quarantine and slaughter. Attempts to vaccinate animals using infected cell extracts, supernatants of infected pig peripheral blood leukocytes, purified and inacti-vated virions, infected glutaraldehyde-fixed macrophages, or detergent-treated in-fected alveolar macrophages have failed to induce protective immunity (4–7). Homol-ogous protective immunity does develop in pigs that survive viral infection. Pigs surviving acute infection with moderately virulent or attenuated variants of ASFV develop long-term resistance to homologous, but rarely to heterologous, virus chal-lenge (8, 9). Pigs immunized with live attenuated ASF viruses containing engineered deletions of specific ASFV virulence-associated genes were protected when challenged with homologous parental virus. Specifically, individual deletion ofUK(DP69R),23-NL (DP71L),TK(A240L), and9GL(B119L) genes or deletion of several genes from MGF360 and -505 (MGF360/505) in the genomes of pathogenic ASF viruses (Malawi Lil-20/1, Pretoriuskop/96/4, E70, and Georgia 2007) markedly attenuated the virus in swine, and the animals immunized with these attenuated viruses were protected against challenge with homologous virus (10–15). These observations constitute the only experimental evidence describing the rational development of an effective live attenuated virus against ASFV.

In particular, deletion of 9GL (B119L) in the highly virulent ASFV isolates Malawi Lil-20/1 and Pretoriuskop/96/4 (Pret4Δ9GL) (10, 16) and deletion of theUKgene in the ASFV E70 isolate (17) resulted in complete attenuation of the viruses in swine even after administration of the recombinant viruses via intramuscular (i.m.) injection at relatively high doses. Interestingly, 9GL gene deletion in highly virulent ASFV Georgia 2007 (ASFV-G-Δ9GL) induced attenuation, but only if inoculated at relatively low doses (13). Therefore, we recently attempted to enhance ASFV-G-Δ9GL attenuation by deleting six genes from MGF360/505, a deletion that by itself was already shown to effectively attenuate the ASFV Georgia 2007 isolate (12). A recombinant virus harboring deletion of the9GLand MGF360/505 genes was attenuated even at relatively high doses (106

HAD50) but showed a decreased ability to replicate in swine and failed to induce

protection against virulent challenge (14). Therefore, we decided to extend our assess-ment of increasing ASFV-G-Δ9GL attenuation by including another virus gene involved in virulence,UK(17). Here, we report the construction of a recombinant ASFV, Δ9GL/ ΔUK virus (Δ9GL/ΔUKv), lacking the9GLandUKgenes, derived from the highly virulent ASFV Georgia 2007 (ASFV-G) isolate. In vivo, ASFV-G-Δ9GL/ΔUK administered i.m. to swine at relatively high doses (106HAD

50) did not induce disease. Importantly, animals

challenged with ASFV-G were protected against the presentation of clinical disease as early as 14 days after inoculation with ASFV-G-Δ9GL/ΔUK. Interestingly, presentation of protection in the animals correlates with the appearance of serum anti-ASFV antibod-ies, but not with circulating virus-specific gamma interferon (IFN-␥)-producing cells.

RESULTS

Development of the 9GL/UK gene deletion mutant of the ASFV-G isolate. ASFV-G-Δ9GL/ΔUK was constructed from the highly pathogenic ASFV-G isolate by a series of two successive homologous-recombination procedures. In the first, a 173-bp

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region, encompassing amino acid residues 11 to 68, within the9GLgene (Fig. 1A) was deleted from ASFV-G and replaced with a cassette containing the p72GUS reporter gene cassette (see Materials and Methods). The recombinant virus was obtained after 11 successive plaque purification events on monolayers of primary swine macrophage cultures (13). The second recombination event replaced theUKgene with a cassette containing the fluorescent mCherry gene under the ASFV p72 promoter. A 255-bp region, encompassing amino acid residues 1 to 85, within theUKgene (Fig. 1B) was deleted from ASFV-G and replaced with a cassette containing the p72mCherry reporter gene cassette (see Materials and Methods). Recombinant virus was selected after 10 rounds of limiting-dilution purification based on the fluorescence activity. The virus population obtained from the last round of purification was amplified in primary swine macrophage cultures to obtain a virus stock.

To ensure the absence of parental ASFV-G, virus DNA was extracted from the virus stock and analyzed by PCR using primers targeting the MGF505-1R and U⌲genes. While amplicons for MGF505-1R genes were detected in DNA extracted from the virus stock, no amplicons were generated with primers targeting the UK gene (data not shown), indicating the lack of contamination of the ASFV-G-Δ9GL/ΔUK stock with parental ASFV-G-Δ9GL.

Analysis of the ASFV-G-9GL/UK full genome sequence relative to the pa-rental ASFV-G sequence.To evaluate the accuracy of the genetic modification and the integrity of the genome of the recombinant virus, full genome sequences of ASFV-G-Δ9GL/ΔUK and parental ASFV-G were obtained using next-generation sequencing (NGS) and compared (Table 1). The DNA sequence assemblies of ASFV-G-Δ9GL/ΔUK and ASFV-G revealed a deletion of 173 nucleotides within the9GLgene corresponding to the introduced modification. The consensus sequence of the ASFV-G-Δ9GL/ΔUK ge-nome showed an insertion of 2,324 nucleotides within the9GLgene corresponding to the p72-␤GUS cassette sequence introduced within the 173-nucleotide deletion in the 9GLgene. In addition, the DNA sequence assemblies of ASFV-G-Δ9GL/ΔUK and ASFV-G revealed a deletion of 255 nucleotides in theUKgene corresponding to the introduced modification. The consensus sequence of the ASFV-G-Δ9GL/ΔUK genome showed an insertion of 937 nucleotides within the UK gene corresponding to the p72mCherry cassette sequence introduced into the 255-nucleotide deletion in theUKgene. Besides the insertion of the cassette, only one additional difference was observed between the ASFV-G-Δ9GL/ΔUK and ASFV-G genomes, a G-to-C nucleotide mutation at position 36465, resulting in an amino acid residue substitution (Glu to Gln) at residue position 224 in the MGF 505-4R open reading frame (ORF). In summary, ASFV-G-Δ9GL/ΔUK did not accumulate any significant mutations during the process of homologous recom-bination and plaque purification (Table 1).

Replication of ASFV-G-9GL/UK in primary swine macrophages.Thein vitro growth characteristics of ASFV-G-Δ9GL/ΔUK were evaluated in primary swine macro-phage cultures, the primary cell targeted by ASFV during infection in swine, and compared to those of parental ASFV-G in multistep growth curves (Fig. 2). Cell cultures were infected at a multiplicity of infection (MOI) of 0.01, and samples were collected at 2, 24, 48, 72, and 96 h postinfection (hpi). Although the three viruses showed similar titers at the end of the experiment, ASFV-G-Δ9GL/ΔUK displayed growth kinetics that were significantly delayed compared to those of ASFV-G-Δ9GL and parental ASFV-G. Δ9GL/Δ9UK yields were approximately 10-fold lower than those of ASFV-G-Δ9GL at 24, 48, and 72 hpi, and depending on the time point considered, were 100- to 1,000-fold lower than those of the parental virus, ASFV-G. Therefore, deletion of theUK gene significantly decreased the ability of ASFV-G-Δ9GL relative to that of the parental ASFV-G isolate to replicatein vitroin primary swine macrophage cultures.

Assessment of ASFV-G-9GL/UK virulence in swine.To evaluate the degree of attenuation reached by ASFV-G-Δ9GL/ΔUK, 80- to 90-pound pigs were inoculated i.m. with either 102, 104, or 106HAD

50and compared with naive animals inoculated with

102 HAD

50 of parental ASFV-G. All the animals inoculated with ASFV-G exhibited

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FIG 1Amino acid sequence alignment of protein products of9GL(B119L) andUK(DP96R) from different ASFV isolates. Virus isolates of various temporal and geographic origins, including those obtained from ticks and pig sources, were compared. The partial deletions introduced into ASFV-G that yielded ASFV-G-Δ9GL/ΔUK are shown between brackets. (.), identical amino acid residues; (*), stop codon.

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increased body temperature (⬎104°F) by day 6 postinfection, presenting with clinical signs associated with the disease, including anorexia, depression, purple skin discolor-ation, staggering gait, and diarrhea (Table 2). Signs of the disease were aggravated progressively over time, and the animals either died or were euthanizedin extremisby days 7 to 9 postinfection. Conversely, animals inoculated i.m. with any of the prescribed doses of ASFV-G-Δ9GL/Δ9UK did not present with any ASF-related signs, remaining clinically normal during the entire observation period (21 days). Therefore, deletion of the9GLandUKgenes produced complete attenuation of the parental virulent ASFV-G. Deletion of theUKgene enhanced attenuation of ASFV-G-Δ9GL to the extent that 106

HAD50of ASFV-G-Δ9GL/ΔUK was completely attenuated, while animals inoculated with

104HAD

50of ASFV-G-Δ9GL presented various levels of disease (13).

Analysis of viremia in animals infected with different doses of ASFV-G-Δ9GL/ΔUK revealed differential patterns depending on the administered dose. Animals receiving 102 HAD

50showed a heterogeneous pattern, with approximately equal numbers of

animals presenting maximum high titers (106to 107HAD

50/ml), moderately high titers

(103to 104HAD

50/ml), and low or undetectable titers (Fig. 3A). Viremia began to be

detected by 7 days postinfection (dpi), peaking by 11 to 14 dpi and decreasing in most

FIG 2In vitro growth characteristics of ASFV-G-Δ9GL/ΔUK and parental ASFV-G-Δ9GL and ASFV-G. Primary swine macrophage cultures were infected (MOI⫽0.01) with each of the viruses, and the virus yields were titrated at the indicated times postinfection. The data represent the means and standard deviations from three independent experiments. The sensitivity of virus detection was ⱖ1.8 log10

[image:5.585.42.371.94.199.2]

HAD50/ml.

TABLE 1Summary of differences between the full-length genome sequence of ASFV-G-Δ9GL and the parental ASFV-G compared with ASFV Georgia07/1

NPNa Type of modification

Virus presencee

ASFV-G ASFV-G9GL/UK

433 T insertion ⫹ ⫹

411 A insertion ⫹ ⫹

1602 MGF 360-1L TT deletion FSb

1620 MGF 360-1L T insertion FS ⫹ ⫹

36465 MGF 505-4R G-to-C Glu224Gln ⫺ ⫹

97391 B438L A-to-G SMc ⫹ ⫹

166192 E199L C-to-G Ala85Pro ⫹ ⫹

183303 T insertion in an NCRd ⫹ ⫹

aNPN, nucleotide position number (based on the sequence of the ASFV Georgia 2007/1 isolate published by

Chapman et al. 3).

bFS, nucleotide modification causes frameshift in the corresponding ORF. cSM, nucleotide modification causes silent mutation.

dNCR, noncoding region. e, present;, absent.

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cases to moderate titers by 21 dpi. On the other hand, most of the animals receiving 104HAD

50showed clear, detectable viremia by 7 dpi, peaking at high titers by 11 dpi,

and decreasing in most cases to moderate titers by 21 dpi (Fig. 3B). Lastly, animals receiving 106 HAD

50in most cases showed high viremia titers that could be clearly

detected by 4 dpi, peaking by 7 to 11 dpi, and decreasing in most cases to moderate titers by 21 dpi (Fig. 3C).

Protective efficacy of ASFV-G-9GL/UK against challenge with parental ASFV-G. A series of three independent experiments were performed in order to assess the ability of ASFV-G-Δ9GL/Δ9UK infection to induce protection against challenge with highly virulent parental ASFV-G. Animals were inoculated i.m. with either 102, 104, or 106

HAD50of ASFV-G-Δ9GL/ΔUK and challenged 28 days later with 103HAD50of ASFV-G by

the i.m. route. Five naive animals that were challenged in each experiment using the same route and dose served as a mock-inoculated/challenge control group.

All the mock-inoculated animals started showing disease-related signs by 3 to 5 days postchallenge (dpc), with rapidly increasing disease severity in the following hours, and were euthanized between 5 and 8 dpc (Table 3). On the other hand, out of nine animals infected with 102HAD

50of ASFV-G-Δ9GL/ΔUK, after challenge, five presented with a

severe form of the disease with kinetics almost indistinguishable from those of the mock-infected and challenged animals. The remaining four animals in the group did not show any significant sign of the disease.

Importantly, all 10 animals infected with 104HAD

50of ASFV-G-Δ9GL/ΔUK remained

completely asymptomatic after challenge throughout the observational period (21 days).

Interestingly, animals infected with 106 HAD

50 of ASFV-G-Δ9GL/ΔUK presented

heterogeneous behavior after being challenged. Considering all three experiments together (Table 3), there were 4 out of 15 animals presenting with a late (10 or 11 dpc), transient rise in body temperature without showing any additional clinical signs related to the disease. Another 10 animals in the group remained completely asymptomatic during the observational period. Finally, one animal (in the third experiment) became sick around 9 dpc, rapidly evolved into severe disease, and was euthanized by 11 dpc. Analysis of viremia after challenge in these animals revealed that, as expected, the levels of viremia corresponded to the presence of severe clinical disease (Fig. 3A, B, and C). Protected animals, regardless of the dose of ASFV-G-Δ9GL/ΔUK received, presented viremia kinetics after challenge that, after a small transient peak, decreased to low (103

HAD50/ml) or undetectable (in 17 out of the total 25 protected animals) levels by 21

dpc. Conversely, in all the animals that succumbed to the challenge, viremia peaked after challenge to values between 107and 108HAD

50/ml.

Detection of ASFV-G-Δ9GL/ΔUK or challenge ASFV-G in the blood of surviving animals was performed by fluorescence microscopy and the presence of theUKgene by PCR (as described in Materials and Methods), respectively. Determinations were performed using the sample at the time postchallenge showing the highest viremia titer. In general, regardless of the dose administered, all surviving animals inoculated with ASFV-G-Δ9GL/ΔUK showed the presence of the virus. Interestingly, these animals also showed circulating ASFV-G in 50%, 30%, and 20% of the animals receiving 102, 104,

or 106HAD

[image:6.585.42.545.85.163.2]

50of ASFV-G-Δ9GL/ΔUK, respectively (data not shown).

TABLE 2Swine survival and fever response following infection with different doses of ASFV-G-Δ9GL/ΔUK or parental ASFV-Ga

Virus Dose (HAD50)

No. of survivors/total

Mean time to death [days (SD)]

Fever Time to onset [days (SD)]

Duration [days (SD)]

Max daily temp [°F (SD)]

ASFV-G 102 0/10 8.6 (1.49) 6 (1.8) 3.2 (0.84) 106.2 (1.38)

ASFV-G-Δ9GL/ΔUK 102 9/9 103.1 (0.46)

ASFV-G-Δ9GL/ΔUK 104 10/10 103.2 (0.32)

ASFV-G-Δ9GL/ΔUK 106 15/15 103.4 (0.59)

aData are combined from the results of three independent experiments. SD, standard deviation.

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Therefore, animals treated with ASFV-G-Δ9GL/ΔUK, administered at the appropriate dose, were protected against the presentation of clinical disease when challenged with the highly virulent parental virus. Nevertheless, protection did not impede replication of challenge virus in a significant proportion of the animals.

FIG 3Viremia titers detected in pigs i.m. inoculated with either 102(A), 104(B), or 106(C) HAD 50of

ASFV-G-Δ9GL/ΔUK and i.m. challenged (indicated by arrows) 28 days later with 103HAD

50of ASFV-G.

Each curve represents data from an individual animal. The gray circles show the viremia of control animals infected with 102HAD

50. The sensitivity of virus detection wasⱖ1.8 log10HAD50/ml.

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Considering the good level of protection against virulent challenge in animals infected 28 days earlier with 104 HAD

50 of ASFV-G-Δ9GL/ΔUK, it was interesting to

assess how early that status was achieved after ASFV-G-Δ9GL/ΔUK infection. Three groups (n⫽5) of animals were i.m. infected with 104HAD

50of ASFV-G-Δ9GL/ΔUK and

challenged 7, 14, or 21 days later with 103HAD

50of ASFV-G by the i.m. route (Table 4).

[image:8.585.40.548.84.219.2]

An additional group were mock vaccinated and served as controls. The mock-vaccinated animals started showing disease-related signs around 3 dpc, with rapid progression in disease severity, followed by euthanasia by 5 dpc. The group of animals challenged at day 7 post-ASFV-G-Δ9GL/ΔUK infection presented a rise in body tem-perature by 3 to 4 dpc. Four of the animals rapidly developed severe disease and were euthanized at 5 dpc. The remaining animal quickly recovered and continued during the rest of the observational period without showing any ASF-related clinical signs. Inter-estingly, all the animals challenged at 14 days post-ASFV-G-Δ9GL/ΔUK infection sur-vived. Three of them remained clinically normal during the 21-day observational period, and the other two showed a transient rise in body temperature around day 11 postchallenge, with no other clinical signs of disease. Finally, four of five animals challenged at 21 days after ASFV-G-Δ9GL/ΔUK infection did not present with any clinical signs of disease during the 21 days of observation. The remaining animal became sick at 15 dpc and was euthanized at 17 dpc.

TABLE 3Swine survival and fever response in ASFV-G-Δ9GL/ΔUK-infected animals challenged with parental ASFV-Ga

Virus Dose (HAD50)

No. of survivors/total

Mean time to death [days (SD)]

Fever No. of days to onset [days (SD)]

[image:8.585.40.545.615.695.2]

Duration [days (SD)]

Maximum daily temp [°F (SD)]

Mock 0/5 7.6 (0.55) 3.6 (0.55) 5 (0.50) 105.6 (0.7)

ASFV-G-Δ9GL/ΔUK 102 2/5 7. (0)b 3.34 (0.68)b 3.7 (0.58)b 106.1 (0.87)b

ASFV-G-Δ9GL/ΔUK 104 5/5 102.9 (1.81)

ASFV-G-Δ9GL/ΔUK 106 5/5 10.5 (4.26)c 4 (2.81)c 104.9 (0.78)c

Mock 0/5 5 (0) 4.8 (0.45) 0.2 (0.45) 106.2 (0.91)

ASFV-G-Δ9GL/ΔUK 102 2/4 5.5 (0.71)d 5 (0)d 1.5 (0.71)d 105.8 (0.7)d

ASFV-G-Δ9GL/ΔUK 104 5/5 103.4 (0.92)

ASFV-G-Δ9GL/ΔUK 106 5/5 10e 3e 104.7e

Mock 0/5 5 (0) 3.8 (0.45) 1.2 (0.45) 105.7 (0.32)

ASFV-G-Δ9GL/ΔUK 106 4/5 11f 11 (2.64)f 3 (1)f 104.5 (0.57)f

aData from three independent experiments are presented. All animals were i.m. challenged at 28 days post-ASFV-G-Δ9GL/ΔUK infection with 103HAD

50of ASFV-G. bData are based on 3 of 5 animals presenting severe disease symptoms in the first two experiments. The remaining two animals in the first and second experiments

did not present with any ASFV-related symptoms, and their average maximum daily body temperature was 103.1°F (SD, 0.85).

cData are based on 2 of 5 animals transitorily presenting a rise in body temperature and very mild disease symptoms. The remaining 3 animals did not present with

any ASFV-related symptoms, and their average maximum daily body temperature was 102.6°F (SD, 0.15).

dData are based on 2 of 4 animals presenting severe disease symptoms in the first two experiments. The remaining two animals in the first and second experiments

did not present with any ASFV-related symptoms, and their average maximum daily body temperature was 103.1°F (SD, 0.85).

eData are based on 1 of 5 animals transitorily presenting a rise in body temperature. The remaining 4 animals did not present with any ASFV-related symptoms, and

their average maximum daily body temperature was 103.2°F (SD, 0.47).

fData are based on 1 of 5 animals transitorily presenting a rise in body temperature and another presenting severe disease symptoms euthanized on day 11

postchallenge. The remaining 3 animals did not present with any ASFV-related symptoms or a rise in body temperature.

TABLE 4Swine survival and fever response in animals challenged with parental ASFV-G at different times post-ASFV-G-Δ9GL/ΔUK infectiona

Virus (104HAD 50)

Time of challenge (dpi)

No. of survivors/total

Mean time to death [days (SD)]

Fever Time to onset [days (SD)]

Duration [days (SD)]

Maximum daily temp [°F (SD)]

Mock 0/5 5 (0) 3.8 (0.45) 1.2 (0.45) 105.7 (0.32)

ASFV-G-Δ9GL/ΔUK 7 1/5 4.75 (1.5) 3.2 (0.45) 1.4 (1.52) 105.8 (1.06)

ASFV-G-Δ9GL/ΔUK 14 5/5 11 (1.41)b 3.5 (3.52)b 105.2 (0.28)b

ASFV-G-Δ9GL/ΔUK 21 4/5 17c 15c 2c 105.8c

aAll animals were i.m. challenged with 103HAD

50of ASFV-G at the indicated time post-ASFV-G-Δ9GL/ΔUK infection.

bData are based on 2 of 5 animals transitorily presenting a rise in body temperature without ASFV-related symptoms. The remaining 3 animals did not present with

any ASFV-related symptoms, and their average maximum daily body temperature was 103.1°F (SD, 0.7).

cData are based on 1 of 5 animals presenting severe disease symptoms. The remaining 4 animals did not present with any ASFV-related symptoms, and their average

maximum daily body temperature was 102.6°F (SD, 0.75).

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Viremia kinetics after challenge in the four animals succumbing to challenge at 7 days post-ASFV-G-Δ9GL/ΔUK infection were similar to those in the mock-treated chal-lenge animals. The surviving animal had viremia equivalent to that of unprotected pigs, remaining at high titers until the end of the experimental period (Fig. 4A). At the time of challenge, the animals in the group challenged at 14 days post-ASFV-G-Δ9GL/ΔUK infection all presented significantly high viremia titers. After challenge, these animals FIG 4Viremia titers detected in pigs i.m. challenged (indicated by arrows) with 103HAD

50of ASFV-G at

7 (A), 14 (B), or 21 (C) days post-i.m. infection with 104HAD

50of ASFV-G-Δ9GL/ΔUK. Each curve represents

data from an individual animal. The circles show the viremias of mock-vaccinated and challenged animals. The sensitivity of virus detection wasⱖ1.8 log10HAD50/ml.

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(all protected) presented extended viremia that remained high in two of them and declined considerably in the other three by the end of the experimental period (Fig. 4B). A very similar scenario was observed in the group challenged at 21 days post-ASFV-G-Δ9GL/ΔUK infection, with declining viremia titers in most of the protected animals by the end of the experiment (Fig. 4C).

Detection of ASFV-G-Δ9GL/ΔUK or challenge ASFV-G in the blood of surviving animals, regardless of the time of challenge, showed the presence of ASFV-G-Δ9GL/ ΔUK. The challenge virus was detected in the only survivor challenged at 7 days post-ASFV-G-Δ9GL/ΔUK infection, in one of the five surviving animals challenged at 14 days post-ASFV-G-Δ9GL/ΔUK infection, and in none of the survivors challenged at 21 days after ASFV-G-Δ9GL/ΔUK infection.

Therefore, ASFV-G-Δ9GL/ΔUK is able to induce protection against the presentation of clinical disease after 14 dpi. This is quite a remarkable fact, since the vast majority of reports on protection induced in animals infected with attenuated ASFV strains present results obtained with challenges performed no earlier than 4 to 6 weeks postinfection. Analysis of the host immune response in animals infected with ASFV-G-9GL/

UK.Host immune mechanisms mediating protection against virulent strains of ASFV in animals infected with attenuated strains of virus are not well identified (18–20). In order to gain additional understanding of immune mechanisms present in animals protected against ASFV, we attempted to correlate the presence of immunological parameters and protection against challenge. We focused on quantifying the presence of anti-ASFV circulating antibodies at the time of challenge, circulating ASFV-specific IFN-␥-producing cells, and systemic levels of several mediators of the innate immune response.

The antibody response against ASFV antigens was monitored at the time of chal-lenge of the ASFV-G-Δ9GL/ΔUK-infected animals. An ASFV-specific antibody response was detected in the sera of these animals, using two in-house-developed assays, a direct enzyme-linked immunosorbent assay (ELISA) and an immunoperoxidase assay (IPA) (14). All five animals infected with 102 HAD

50 of ASFV-G-Δ9GL/ΔUK that were

unprotected when challenged 28 days later did not have measurable anti-ASFV titers by ELISA, and only two of them were barely positive by IPA. In contrast, four of the five animals surviving the challenge possessed circulating anti-ASFV antibodies, although some of them were at low levels (Fig. 5A). A clearer scenario was found in the animals infected with 104 HAD

50of ASFV-G-Δ9GL/ΔUK and challenged 28 days later. All the

animals in this group were protected, and in all cases, the animals possessed anti-ASFV antibodies at the moment of challenge (Fig. 5B). Antibody titers in these animals were generally higher than in those inoculated with 102HAD

50of ASFV-G-Δ9GL/ΔUK. All 14

protected animals infected with 106 HAD

50of ASFV-G-Δ9GL/ΔUK and challenged 28

days later presented high anti-ASFV antibody titers (Fig. 5C). The only unprotected animal in the group showed low antibody titers (10- to 100-fold lower than those of the groupmates).

Analysis of the antibody response in animals infected with 104 HAD

50of

ASFV-G-Δ9GL/ΔUK and challenged at different times postinfection also supports a correlation between the presence of anti-ASFV antibodies and protection. ASFV-specific antibodies were undetectable in all five animals challenged at 7 days post-ASFV-G-Δ9GL/ΔUK infection (all but one were unprotected against the challenge) (Fig. 6A). Conversely, in all the animals challenged at 14 days post-ASFV-G-Δ9GL/ΔUK infection, it was possible to detect significant anti-ASFV antibody titers (Fig. 6B). Animals challenged at 21 days post-ASFV-G-Δ9GL/ΔUK infection presented with significant anti-ASFV antibody titers (Fig. 6C).

As a marker of T-cell sensitization, the ASFV-specific IFN-␥response, detected by enzyme-linked immunosorbent spot (ELISpot) assay, was evaluated at the time of challenge with the virulent parental ASFV-G. Analysis of animals inoculated with 102

HAD50and 104HAD50of ASFV-G-Δ9GL/ΔUK was performed in the second experiment

and in animals inoculated with 106HAD

50of ASFV-G-Δ9GL/ΔUK in the second and third

experiments (Table 2). There were greater numbers of ASFV-specific T cells producing

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IFN-␥in surviving animals infected with 102HAD

50of ASFV-G-Δ9GL/ΔUK than in those

that had to be euthanized due to the presence of severe disease (Fig. 5A). All the animals infected with 104HAD

50of ASFV-G-Δ9GL/ΔUK (which were protected)

exhib-ited high levels of ASFV-specific T cells producing IFN-␥(Fig. 5B). All 10 animals infected with 106HAD

50of ASFV-G-Δ9GL/ΔUK had high levels of ASFV-specific T cells producing

IFN-␥, including the animal that did not survive the challenge (Fig. 5C). Analysis of the presence of ASFV-specific T cells producing IFN-␥in animals infected with 104HAD

50

of ASFV-G-Δ9GL/ΔUK and challenged at 7 days showed no differences between pro-tected and unpropro-tected individuals (Fig. 6A). All the propro-tected animals challenged at 14 days postinfection with ASFV-G-Δ9GL/ΔUK displayed heterogeneous levels of cells producing IFN-␥(Fig. 6B), and the only animal that succumbed when challenged at 21 days post-ASFV-G-Δ9GL/ΔUK infection had the lowest number of cells producing IFN-␥ in that group (Fig. 6C). Therefore, there was no direct association in ASFV-G-Δ9GL/ΔUK-FIG 5Serological ASFV-specific antibodies detected by ELISA and immunoperoxidase staining (IPA), along with the number of circulating ASFV-specific IFN-␥-producing cells for each individual pig at the time of challenge in animals infected with either 102(A), 104(B), or 106(C) HAD

50of ASFV-G-Δ9GL/ΔUK. The survival status of the swine is

indicated as survival (open symbols) or no survival (solid symbols). neg, negative; dotted lines, threshold for positivity.

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FIG 6Serological ASFV-specific antibodies detected by ELISA and IPA, along with the number of circulating ASFV-specific IFN-␥-producing cells, for each individual pig at the time of challenge at 7 (A), 14 (B), or 21 (C) days postinfection with 104HAD

50of ASFV-G-Δ9GL/ΔUK. The survival status of the swine

is indicated as survival (open symbols) or no survival (solid symbols). Dotted lines, threshold for positivity.

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infected animals between the development of virus-specific IFN-␥-producing cells and resistance to challenge with virulent ASFV-G.

Protection against virulent challenge in animals infected with attenuated ASFV strains has not been previously reported at such early times as described here. Therefore, we tried to establish an association between a specific pattern of innate host immune response mediators and protection against challenge in ASFV-G-Δ9GL/ΔUK-infected animals, particularly at early times postinfection. Levels of MCP2, transforming growth factor beta (TGF-␤), IFN-␣, IFN-␤, IFN-␥, interleukin 1␣(IL-1␣), IL-1␤, IL-2, IL-5, IL-6, IL-8, IL-10, IL-12-p35, IL-12-p40, OAS, PKR, tumor necrosis factor (TNF), MX-1, and VCAM in sera of animals challenged at 7 and 14 days post-ASFV-G-Δ9GL/ΔUK infection were assessed using commercial ELISAs following the manufacturers’ protocols. The results demonstrated that, in general, the circulating levels of all the detected cytokines were heterogeneous in every group considered (Fig. 7). Regardless of heterogeneity, in general, average values did not vary significantly between protected and nonprotected animals for most of the cytokines assessed, and in general, the values did not signifi-cantly differ from those of naive animals. In contrast to this general observation for most of the cytokines, systemic IFN-␣and TGF-␤levels were elevated in the only animal surviving challenge at 7 days post-ASFV-G-Δ9GL/ΔUK infection relative to the other animals in the group, as well as the mock-treated animals. Similarly, in all the animals challenged at 14 days post-ASFV-G-Δ9GL/ΔUK infection, it was possible to detect high levels of IFN-␣and TGF-␤.

DISCUSSION

The use of attenuated strains is currently the most plausible approach to develop an effective ASF vaccine. Rational development of attenuated strains by genetic manipu-lation is a valid alternative to, and perhaps safer methodology than, the use of naturally attenuated isolates (8, 21). Several attenuated strains, obtained by genetic manipula-tion consisting of single-gene delemanipula-tions, have been shown to induce protecmanipula-tion against the virulent parental virus (10–13, 15, 17). In some cases, a single-gene deletion did not completely attenuate the virus, particularly when it was attempted to increase immu-nogenicity by using larger quantities of virus (13). We hypothesized that combined deletion of virulence-associated genes would modulate the resulting recombinant virus. Until recently, this type of genetic manipulation in field isolates (i.e., isolates not adapted to replication in any cell line) had not been reported. Recently, we developed ASFV-G-Δ9GL/ΔMGF (14), a recombinant derivative of the ASFV Georgia 2007 isolate harboring all the gene deletions individually present in two other attenuated deriva-tives, ASFV-G-Δ9GL (13) and ASFV-G-ΔMGF (12). ASFV-G-Δ9GL/ΔMGF presented a significantly more attenuated phenotype than ASFV-G-Δ9GL, which showed remaining virulence when used at relatively high doses and the inconvenience of decreased replication during infection in swine, losing all immunogenicity and protective effect (14).

Here, it is shown that an additional deletion of the virulence-associated UKgene effectively augments the attenuation of the9GLgene deletion. ASFV-G-Δ9GL/ΔUK was completely attenuated even at doses 100 times higher than those at which ASFV-G-Δ9GL became virulent (104HAD

50) (13).

ASFV-G-Δ9GL/ΔUK effectively protected 100% of the animals with a dose of 104

HAD50when challenged at 28 dpi, while a dose of 102HAD50protected only 40% of the

animals. This is interesting, because at the lower dose, ASFV-G-Δ9GL was able to protect all the animals (13), suggesting that the additional deletion of theUKgene decreased the protective effect of ASFV-G-Δ9GL/ΔUK, perhaps because it had a decreased repli-cation rate in swine macrophages compared to ASFV-G-Δ9GL. This is in agreement with the lower replication rate of Δ9GL/ΔUK compared with its parental ASFV-G-Δ9GL in primary cultures of swine macrophages. It is possible that ASFV-G-ASFV-G-Δ9GL/ΔUK also replicates less efficiently in swine, inducing a less effective protective host response. It is also possible that either the reporter proteins or the use of the p72 promoter for both ␤-glucuronidase (GUS) and mCherry could possibly negatively affect ASFV virulence or

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FIG 7Evaluation of systemic levels of different host cytokines at the time of challenge, either 7 or 14 days postinfection with ASFV-G-Δ9GL/ ΔUK. The values were determined for individual animals and are expressed as the concentration per milliliter of serum, as described in Materials and Methods. The survival status of the swine is indicated as survival (open symbols) or no survival (solid symbols).

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could compete with the transcription of p72. In ASFV-G-Δ9GL/ΔUK, we did not see any decrease in virus replication compared to ASFV-G-Δ9GL, suggesting that using the p72 promoter for both reporter genes did not negatively affect virus replication.

Higher doses of ASFV-G-Δ9GL/ΔUK (106HAD

50) appear not to be as effective as 104

HAD50, since some of the animals (4 out of 15) presented a transient rise in body

temperature, and 1 animal developed serious ASF disease and needed to be eutha-nized, while none of the animals receiving 104HAD

50were clinically affected after the

challenge.

Interestingly, ASFV-G-Δ9GL/ΔUK (used at a dose of 104HAD

50) was able to induce

a significant level of protection against challenge as early as 14 dpi. These data, along with those obtained with the attenuated virus strain Pret4Δ9GL (unpublished data), are the first reports demonstrating the presence of early protection achieved in animals infected with attenuated strains and then challenged with homologous ASFV. At this moment, it is not clear what host mechanisms mediate this early protection. In fact, immune mechanisms mediating protection against ASFV infection and disease, partic-ularly in animals infected with attenuated strains, remain largely uncharacterized.

The only direct evidence regarding mechanisms of protection against ASFV is derived from reports in which acquisition of humoral protective immunity to ASFV by naive pigs was achieved after passive transfer of ASFV-specific antibodies obtained from donor pigs that were previously infected with attenuated ASFV (18, 21). The presence of circulating virus-neutralizing antibodies in animals infected with attenu-ated ASFV strains has been a controversial issue (16, 18, 20–22). Moreover, there have been contradictory reports regarding the efficacy of neutralizing antibodies in prevent-ing infection or disease (16, 22). The only direct evidence related to cellular immunity is derived from a report demonstrating that partial CD8⫹T-cell depletion impedes the development of a host protective immune response in challenged animals that have been previously vaccinated with attenuated ASFV (19).

The presence of anti-ASFV specific antibodies appears to be associated with pro-tection. Absent at 7 days post-ASFV-G-Δ9GL/ΔUK infection, when almost all animals succumbed to challenge, they were clearly present at 14 dpi and onward, when most of the animals survived challenge. Interestingly, nonprotected animals pretreated with 102HAD

50of ASFV-G-Δ9GL/ΔUK did not have detectable levels of virus-specific

anti-bodies, and the only animal that succumbed to challenge among those treated with 106 HAD

50of ASFV-G-Δ9GL/ΔUK had the lowest antibody titer in that group. These

results are in agreement with results obtained in studies using the Pret4Δ9GL virus (unpublished data).

Recently, a possible correlation between the induction of virus-specific circulating IFN-␥-producing cells and protection in animal vaccinated with attenuated ASFV strains (23) or with experimental DNA vaccines has been reported (24). Our results indicated no close association between virus-specific IFN-␥-producing cells and protection as an independent factor. As an example, all four animals succumbing to the challenge at 7 days post-ASFV-G-Δ9GL/ΔUK infection had significant levels of IFN-␥-producing cells and complete absence of virus-specific antibodies. It is not clear at this point what may account for these differences.

A note of interest is that in all groups all the protected animals had the highest viremia values after infection with ASFV-G-Δ9GL/ΔUK. Even in the group of animals challenged at 7 days post-ASFV-G-Δ9GL/ΔUK infection, the protected animals pre-sented with the highest prechallenge viremia. It may be that the highest viremias are more effective in inducing protective host innate immune mechanisms. Although the role of cytokines in ASF pathogenesis has received abundant attention (25–29), the possible role of the innate response in protection is not well understood. Our results indicate that systemic IFN-␣ and TGF-␤ levels were elevated in the only animal surviving challenge at 7 days post-ASFV-G-Δ9GL/ΔUK infection relative to the other animals in the group succumbing to the challenge, as well as the mock-treated ones. Similarly, in all the animals challenged at 14 days post-ASFV-G-Δ9GL/ΔUK infection, it was possible to detect significant levels of IFN-␣ and TGF-␤. Recently, it has been

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reported that MGF genes play a role in blocking the IFN-␣effectin vivo(30), suggesting a relationship between lack of MGF genes in attenuated ASFV strains and the inability of these viruses to block the host IFN-␣response. The direct effect of IFN in inhibiting ASFV growth in vitro is controversial (30–32). Further studies are necessary to fully evaluate the role of IFN-␣and TGF-␤in the prevention of ASFV infection in swine to support the preliminary observation reported here.

In summary, we present here the development of a double-deletion recombinant ASFV that had an enhanced attenuated phenotype relative to its parental ASFV-G-Δ9GL, effectively protecting pigs from lethal challenge. ASFV-G-Δ9GL/ΔUK is the first attenuated virus that has been shown to induce protection against virulent challenge at early times postimmunization and, along with the rationally designed experimental vaccine viruses ASFV-G-Δ9GL (13) and ASFV-G-ΔMGF (12), is the only strain reported to protect against the highly virulent epidemiologically significant ASFV-G isolate.

MATERIALS AND METHODS

Cell cultures and viruses.Primary swine macrophage cultures were prepared from defibrinated swine blood as previously described (15). Briefly, heparin-treated swine blood was incubated at 37°C for 1 h to allow sedimentation of the erythrocyte fraction. Mononuclear leukocytes were separated by flotation over a Ficoll-Paque (Pharmacia, Piscataway, NJ) density gradient (specific gravity, 1.079). The monocyte/macrophage fraction was cultured in plastic Primaria (Falcon; Becton Dickinson Labware, Franklin Lakes, NJ) tissue culture flasks containing macrophage medium composed of RPMI 1640 medium (Life Technologies, Grand Island, NY) with 30% L929 supernatant and 20% fetal bovine serum (HI-FBS; Thermo Scientific, Waltham, MA) for 48 h at 37°C under 5% CO2. Adherent cells were detached

from the plastic by using 10 mM EDTA in phosphate-buffered saline (PBS) and were then reseeded into Primaria T25 6- or 96-well dishes at a density of 5⫻106cells per ml for use in assays 24 h later.

Comparative growth curves between ASFV-G, ASFV-G-Δ9GL, and ASFV-G-Δ9GL/ΔUK were performed in primary swine macrophage cultures. Preformed monolayers were prepared in 24-well plates and infected at an MOI of 0.01 (based on the HAD50previously determined in primary swine macrophage

cultures). After 1 h of adsorption at 37°C under 5% CO2, the inoculum was removed and the cells were

rinsed two times with PBS. The monolayers were then rinsed with macrophage medium and incubated for 2, 24, 48, 72, and 96 h at 37°C under 5% CO2. At appropriate times postinfection, the cells were frozen

at less than⫺70°C, and the thawed lysates were used to determine titers (in HAD50/ml) in primary swine

macrophage cultures. All the samples were run simultaneously to avoid interassay variability. Virus titration was performed on primary swine macrophage cultures in 96-well plates. Virus dilutions and cultures were performed using macrophage medium. The presence of virus was assessed by hemadsorption (HA), and virus titers were calculated by the Reed and Muench method (33).

ASFV-G was a field isolate kindly provided by Nino Vepkhvadze, from the Laboratory of the Ministry of Agriculture (LMA) in Tbilisi, Republic of Georgia.

Construction of the recombinant ASFV-G9GL/UK.Recombinant ASFVs were generated by sequential homologous recombination between the parental ASFV genome and recombination transfer vectors in infection and transfection procedures using swine macrophage cultures (15). First, a recom-binant transfer vector (p72GUSΔ9GL) containing flanking genomic regions, including portions of9GL mapping to the left (1.2 kbp) and right (1.15 kbp) of the gene and a reporter gene cassette containing the GUS gene with the ASFV p72 late gene promoter, p72GUS, was used. The construction created a 173-nucleotide deletion in the9GLORF (amino acid residues 11 to 68) (Fig. 1). The recombinant transfer vector p72GUSΔ9GL was obtained by DNA synthesis (GenScript, Piscataway, NJ, USA). Macrophage cultures were infected with ASFV-G and transfected with p72GUSΔ9GL. Recombinant viruses represent-ing independent primary plaques were purified to homogeneity by successive rounds of plaque assay purification. The intermediate recombinant virus produced, ASFV-G-Δ9GL (13), was then used as a parental virus in infection/transfection procedures using a recombinant transfer vector that would result in deletion of theUKgene from the virus genome. The recombinant transfer vector (p72mCherryΔUK) containing flanking genomic regions of theUKgene mapping to the left (1.156 kbp) and right (1.190 kbp) of the gene and a reporter gene cassette containing the mCherry gene with the ASFV p72 late gene promoter, p72mCherry, was used. This construction created a 255-nucleotide deletion in theUKORF (amino acid residues 1 to 85) (Fig. 1). The recombinant transfer vector p72mCheryΔUK was obtained by DNA synthesis (GenScript, Piscataway, NJ, USA).

PCR.The purity of ASFV-G-Δ9GL/ΔUK in the virus stock was assessed by PCR. Detection of the MGF360/MGF505 genes (as a control for the presence of virus DNA) was performed using the following pair of primers: forward, 5=GAGGATGATTTGCCCTTCACTCA3=; reverse, 5=CGCCACTAGTAAACATTGTTC TATCT3=. These primers amplified a 422-bp fragment of the MGF505-1R gene. Detection of theUKgene was performed using the following pair of primers that amplified a 210-bp fragment of the gene: forward, 5=GTTGTCGTGGATAATGCACC3=; reverse, 5=GGATGGAGCGCATTAGGGAT3=.

NGS of ASFV genomes.ASFV DNA was extracted from infected cells and quantified as described previously (34). Full-length sequencing of the virus genome was performed as described previously (34). Briefly, 1␮g of virus DNA was enzymatically sheared, and the resulting fragmented-DNA size distribution was assessed. Adapters and library barcodes were ligated to the fragmented DNA. The appropriate size range of the adapter-ligated library was collected using the Pippin Prep system (Sage Science), followed

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by normalization of the library concentration. The DNA library was then clonally amplified onto intermediate source plates (ISPs) and enriched. Enriched template ISPs were prepared and loaded onto Ion chips (Thermo Fisher, Waltham, MA) for sequencing. Sequence analysis was performed using Galaxy (https://usegalaxy.org/) and CLC Genomics Workbench (CLCBio).

Animal experiments.Animal experiments were performed under biosafety level 3 conditions in the animal facilities at Plum Island Animal Disease Center (PIADC), Greenport, NY, USA, following a protocol approved by the Institutional Animal Care and Use Committee.

ASFV-G-Δ9GL/ΔUK was assessed for its virulence phenotype relative to the virulent parental ASFV-G using 80- to 90-pound commercial-breed swine. Groups of pigs (n⫽5) were inoculated i.m. either with 102, 104, or 106HAD

50of ASFV-G Δ9GL/ΔUK or with 104HAD50of ASFV-G. Clinical signs (anorexia,

depression, fever, purple skin discoloration, staggering gait, diarrhea, and cough) and changes in body temperature were recorded daily throughout the experiment. In protection experiments, animals were i.m. inoculated with 102, 104, or 106HAD

50of ASFV-G-Δ9GL/ΔUK and 28 days later i.m. challenged with

103HAD

50of the parental virulent ASFV strain Georgia 2007. The presence of clinical signs associated

with disease was detected as described previously. Additional protective experiments were performed with animals i.m. inoculated with 104HAD

50of ASFV-G-Δ9GL/ΔUK and i.m. challenged 7, 14, or 21 days

later with 103HAD

50of the parental virulent ASFV Georgia 2007 strain.

Detection of ASFV-specific antibodies. Anti-ASFV antibodies in sera of infected animals were quantified using two in-house-developed assays. The first was an immunoperoxidase assay in which Vero cells were infected (MOI⫽0.1) with the Vero cell-adapted ASFV Pret4 strain in 96-well plates. The infected cells were fixed in 50% acetone and 50% methanol for 10 min. The plates were blocked with 5% skim milk (Millipore, Billerica, MA) and 0.05% Tween 20 (Sigma, St. Louis, MO) for 1 h at 37°C. Twofold dilutions of the sera were incubated for 1 h at 37°C on the 96-well ASFV-infected Vero cell monolayer. After washing with 1⫻PBS, the presence of anti-ASFV antibodies was detected by using a commercial biotinylated anti-swine IgG, Vectastain avidin-biotinylated enzyme complex, and Vector VIP horseradish peroxidase (Vector Laboratories, CA). Titers were expressed as the inverse log10value of the highest

serum dilution reacting with the infected cells.

The second methodology was an indirect ELISA. Antigen preparation and the ELISA procedure were based on previously described methods (35) with minor adjustments. Briefly, Vero cells were infected with an ASFV adapted to replicate in Vero cells (34) until the cytopathic effect reached 100%. The infected cells were resuspended in water containing protease inhibitor (Roche, New York, NY), followed by the addition of Tween 80 (G-Biosciences, Saint Louis, MO) and sodium deoxycholate (Sigma, Saint Louis, MO) to a final concentration of 1% (vol/vol). Uninfected Vero cells were treated in the same manner, and the antigens were stored at less than or equal to⫺70°C. Maxisorb ELISA plates (Nunc, Saint Louis, MO) were coated with 1␮g per well of either infected-cell or uninfected-cell antigen. The plates were blocked with phosphate-buffered saline containing 10% skim milk (Merck, Kenilworth, NJ) and 5% normal goat serum (Sigma). Each swine serum was tested at multiple dilutions against both infected- and uninfected-cell antigens. ASFV-specific antibodies in the swine sera were detected by an anti-swine IgG-horseradish peroxidase conjugate (KPL, Gaithersburg, MD) and SureBlue Reserve peroxidase sub-strate (KPL). The plates were read at an optical density at 630 nm (OD630) in an ELx808 plate reader

(BioTek, Shoreline, WA). Swine sera were considered positive for ASFV-specific antibodies if the ratio of the OD630of the reaction mixture against infected-cell antigen to that of the reaction mixture against

uninfected-cell antigen was higher than 2.2.

Detection of ASFV-specific IFN--producing cells.Detection of ASFV-specific IFN-␥-producing cells was performed using a modification of the ELISpot porcine IFN-␥method (R&D, Minneapolis, MN). Peripheral blood mononuclear cells (PBMCs) were isolated from 15 ml of porcine blood by Ficoll-Paque Plus gradient (density, 1.077) and washed twice with 1⫻PBS at room temperature. The cell counts were adjusted to 5⫻ 106cells/ml and seeded into 96-well plates. After seeding, the cells were stimulated with a buffer containing

25 ng/ml of phorbol myristate acetate (PMA) and 25 ng/ml of calcium ionomycin or were exposed to ASFV-G at an MOI of 0.5. The cells and stimulators were then immediately transferred to ELISpot plates (as provided in the kit) and incubated for 18 h at 37°C. The steps for washing, as well as for using the detection antibody, streptavidin-allophycocyanin (AP), and BCIP-nitroblue tetrazolium (NBT) chromogen were sequentially per-formed as recommended by the kit’s manufacturer. Reading was perper-formed with an ELISpot plate reader (Immunospot; Cellular Technology Limited) with the following settings: counting mask size, 100%; normalize counts of mask, off; sensitivity, 130; minimum spot size, 0.086 mm2; maximum spot size, 0.2596 mm2. Oversize

spots were estimated at a spot separation of 1; diffuseness, large; and background balance of 67. Cell counts were expressed as the number of spots per 5⫻106PBMCs/ml.

ACKNOWLEDGMENTS

We thank the Plum Island Animal Disease Center Animal Resource Branch for excellent technical assistance. We particularly thank Melanie Prarat for editing the manuscript.

This project was partially funded through the State of Kansas National Bio- and Agro-Defense Facility Fund (NBAF) and the Institute for Infectious Animal Diseases (IIAD) Career Development Program and an interagency agreement with the Science and Technology Directorate of the U.S. Department of Homeland Security under award numbers HSHQDC-11-X-00077 and HSHQPM-12-X-00005. We also thank ARS/USDA-University of Connecticut SCA 58-1940-1-190 for partially supporting the work.

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on November 7, 2019 by guest

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Figure

FIG 1 Amino acid sequence alignment of protein products of 9GL (B119L) and UK (DP96R) from different ASFV isolates
TABLE 1 Summary of differences between the full-length genome sequence of ASFV-G-Δ9GL and the parental ASFV-G compared with ASFV Georgia07/1
TABLE 2 Swine survival and fever response following infection with different doses of ASFV-G-Δ9GL/ΔUK or parental ASFV-Ga
FIG 3 Viremia titers detected in pigs i.m. inoculated with either 10Each curve represents data from an individual animal
+6

References

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