This article was downloaded by: [134.117.10.200]On: 30 September 2013, At: 06:00Publisher: Taylor & FrancisInforma Ltd Registered in England and Wales Registered Number: 1072954 Registered office: MortimerHouse, 37-41 Mortimer Street, London W1T 3JH, UK

Avian PathologyPublication details, including instructions for authors and subscription information:http://www.tandfonline.com/loi/cavp20

Effects of chicken anaemia virus and infectiousbursal disease virus-induced immunodeficiency oninfectious bronchitis virus replication and genotypicdriftRodrigo A. Gallardo a , Vicky L. van Santen a & Haroldo Toro aa Department of Pathobiology, College of Veterinary Medicine, Auburn University,Auburn, AL, 36849, USAAccepted author version posted online: 19 Jun 2012.Published online: 17 Aug 2012.

To cite this article: Rodrigo A. Gallardo , Vicky L. van Santen & Haroldo Toro (2012) Effects of chicken anaemia virus andinfectious bursal disease virus-induced immunodeficiency on infectious bronchitis virus replication and genotypic drift,Avian Pathology, 41:5, 451-458, DOI: 10.1080/03079457.2012.702889

To link to this article: http://dx.doi.org/10.1080/03079457.2012.702889

PLEASE SCROLL DOWN FOR ARTICLE

Taylor & Francis makes every effort to ensure the accuracy of all the information (the “Content”) containedin the publications on our platform. However, Taylor & Francis, our agents, and our licensors make norepresentations or warranties whatsoever as to the accuracy, completeness, or suitability for any purpose ofthe Content. Any opinions and views expressed in this publication are the opinions and views of the authors,and are not the views of or endorsed by Taylor & Francis. The accuracy of the Content should not be reliedupon and should be independently verified with primary sources of information. Taylor and Francis shallnot be liable for any losses, actions, claims, proceedings, demands, costs, expenses, damages, and otherliabilities whatsoever or howsoever caused arising directly or indirectly in connection with, in relation to orarising out of the use of the Content.

This article may be used for research, teaching, and private study purposes. Any substantial or systematicreproduction, redistribution, reselling, loan, sub-licensing, systematic supply, or distribution in anyform to anyone is expressly forbidden. Terms & Conditions of access and use can be found at http://www.tandfonline.com/page/terms-and-conditions

Effects of chicken anaemia virus and infectious bursaldisease virus-induced immunodeficiency on infectiousbronchitis virus replication and genotypic drift

Rodrigo A. Gallardo, Vicky L. van Santen and Haroldo Toro*

Department of Pathobiology, College of Veterinary Medicine, Auburn University, Auburn, AL 36849, USA

We followed changes in a portion of the S1 gene sequence of the dominant populations of an infectiousbronchitis virus (IBV) Arkansas (Ark) vaccine strain during serial passage in chickens infected with theimmunosuppressive chicken anaemia virus (CAV) and/or infectious bursal disease virus (IBDV) as well as inimmunocompetent chickens. The IBV-Ark vaccine was applied ocularly and tears were collected frominfected chickens for subsequent ocular inoculation in later passages. The experiment was performed twice.In both experiments the dominant S1 genotype of the vaccine strain was rapidly and negatively selected in allchicken groups (CAV, IBDV, CAV�IBDV and immunocompetent). Based on the S1 genotype, the sameIBV subpopulations previously reported in immunocompetent chickens and named component (C) 1 to C5emerged both in immunocompetent and immunodeficient chickens. During the first passage differentsubpopulations emerged, followed by the establishment of one or two predominant populations after furtherpassages. Only when the subpopulation designated C2 became established in either CAV-infected or IBDV-infected chickens was IBV maintained for more than four passages. These results indicate that selection doesnot cease in immunodeficient chickens and that phenotype C2 may show a distinct adaptation to thisenvironment. Subpopulations C1 or C4 initially became established in immunocompetent birds but becameextinct after only a few succeeding passages. A similar result was observed in chickens co-infected withCAV�IBDV. These results suggest that the generation of genetic diversity in IBV is constrained. Thisfinding constitutes further evidence for phenotypic drift occurring mainly as a result of selection.

Introduction

Infectious bronchitis virus (IBV) is a single-strandedRNA group 3 coronavirus that shows extensivegenotypic and phenotypic variability. Genetic diversityis generated during viral replication as a result of thelack of proofreading activity of the RNA polymeraseas well as recombination events (Kusters et al., 1987,1990).

The IBV spike (S) protein is responsible for viralattachment to the host cell and is a relevant target forspecific humoral and cellular host immune responses(Cavanagh, 1981, 1983, 1984; Mockett et al., 1984;Cavanagh & Davis, 1986; Koch et al., 1990; Koch &Kant, 1990; Collisson et al., 2000). The bulb end ofthe S protein (S1) exhibits extensive variation amongIBV populations (Kusters et al., 1987, 1989), providinga successful adaptation favouring immunological es-cape. The phenotype of S1 thus represents an im-portant feature of the evolutionary success of thisvirus. Based on S1 gene sequencing, we and othershave previously shown that the dominant genotype/phenotype of IBV ArkDPI-derived vaccine strainschanges during host invasion (McKinley et al., 2008;van Santen & Toro, 2008; Gallardo et al., 2010). Thisis probably due to selective pressure exerted by the

microenvironments of the host tissues on the replicat-ing virus population.

Chickens in production environments are exposed tomultiple stressors and infectious diseases, which impairinnate and adaptive immunity (Hoerr, 2010). Infectiousbursal disease virus (IBDV), chicken anaemia virus(CAV), and Marek’s disease virus are examples ofendemic pathogens altering the host’s humoral andcell-mediated immune function. We have shown thatIBV is frequently found in association with immuno-suppression as indicated by lymphocytic depletion ofboth the thymus and the bursa of Fabricius (Toro et al.,2006). IBV is thus frequently replicating in immuno-deficient birds in commercial settings. According toKilbourne (1994, p. 262): ‘‘in those experiments ofnature in which the immune response is effectivelyremoved as a selective host force, viral evolution doesnot cease but rather appears to falter or follow differentpathways’’. We thus hypothesized that IBV passage inimmunodeficient chicken populations results in distinctevolutionary pathways. To explore this hypothesis weevaluated IBV genotypic drift during continued IBVpassages in immunocompetent chickens or in chickenspreviously inoculated with CAV and/or IBDV.

*To whom correspondence should be addressed: Tel: �1 334 8442662. E-mail: torohar@auburn.edu

Avian Pathology (October 2012) 41(5), 451�458

Received 16 March 2012

ISSN 0307-9457 (print)/ISSN 1465-3338 (online)/12/050451-08 # 2012 Houghton Trust Ltdhttp://dx.doi.org/10.1080/03079457.2012.702889

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13

Materials and Methods

Viruses. A commercially available monovalent live-attenuated ArkDPI-

derived IBV vaccine strain (Intervet, Millsboro, Delaware, USA) was

used in both experiments. The lyophilized vaccine strain was recon-

stituted in sterile tryptose broth containing a commercially available

antibiotic�antimycotic combination (Invitrogen, Carlsbad, California,

USA) and used in serial chicken passages. The vaccine was character-

ized by S1 gene sequencing prior to use. Results for the vaccine lot used

in Experiment 1 have been reported previously (vaccine B; van Santen &

Toro, 2008), and identical results were obtained for the vaccine lot used

in Experiment 2. As reported for the vaccine lot used in Experiment 1,

vaccine subpopulations component (C) 1 to C5 are detectable only

by reverse transcription polymerase chain reaction (RT-PCR) using

specific primers (van Santen & Toro, 2008; further discussed in

Gallardo et al., 2010) and/or by selection of these vaccine subpopula-

tions in individual chickens (Gallardo et al., 2010). Selection of the

same subpopulation in multiple chickens is strong evidence that the

specific subpopulation pre-existed in the vaccine. Thus, prior to use in

the present experiment, the vaccine lot used for Experiment 2 was

inoculated into chickens and the S1 gene sequence of the IBV found in

samples collected from individual chickens was determined. The same

five subpopulations that had been found in individual chickens

inoculated with the vaccine lot used for Experiment 1 were found in

chickens inoculated with the second vaccine lot (van Santen, 2010).

A previously described CAV strain 03-4876 (van Santen et al., 2004)

was used to cause T-lymphocyte deficiency. CAV was propagated and

titrated in MDCC-MSB1 cells by standard procedures (McNulty &

Todd, 2008). The virus stock was aliquoted and stored at �808C. An

aliquot was thawed and evaluated for its efficacy to cause immunode-

ficiency at the dose used in the experiments (100 ml containing 1.6�105

median tissue culture infectious doses) by inoculation into specific

pathogen free chickens. Ten days after intramuscular inoculation,

chickens showed reduced haematocrit levels and severe thymus atrophy.

Histomorphometry analysis performed as previously described (van

Santen et al., 2004) revealed severe thymic lymphocyte depletion.

Aliquots of this virus stock were used for inoculation of subsequent

groups.

The previously characterized variant IBDV AL2 strain (Toro et al.,

2009) was used to induce B-lymphocyte deficiency. The AL2 strain was

propagated, titrated, and stored at �808C. An aliquot of the virus

stock was thawed and tested for its efficacy to cause disease in specific

pathogen free chickens. Five days after inoculation with the dose used in

the experiments (200 ml containing 2�105 median chicken infectious

doses), infected chickens showed reduced bursa weight and reduced

bursa indices, as well as bursa lymphocytic depletion as determined by

histomorphometry. Aliquots of this virus stock were used for inocula-

tion of subsequent groups.

Chickens. White Leghorn chickens hatched from specific pathogen free

fertile eggs (Sunrise Farms, Catskill, New York, USA) were used in all

experiments. Hatched chickens were maintained in isolation in biosafety

level 2 facilities. Experimental procedures and animal care were

performed in compliance with all applicable federal and institutional

animal use guidelines. Auburn University College of Veterinary

Medicine is an Association for Assessment and Accreditation of

Laboratory Animal Care-accredited Institution.

Experimental design. Experiment 1. Four groups each of 10 chickens

were hatched at 8-day intervals for nine serial passages of the IBV

vaccine. On day 7 post hatch, three groups, designated CAV, IBDV and

CAV� IBDV were inoculated intramuscularly with CAV (100 ml

containing 1.6�105 median tissue culture infectious doses), subcuta-

neously with IBDV (200 ml containing 2�105 median chicken infectious

doses) or with both CAV and IBDV, respectively. The fourth group

consisted of uninoculated immunocompetent chickens. For the first

passage of IBV, five extra chickens were added to each group and the

effects of the immunosuppressive viruses were confirmed at day 10 of

age in the extra chickens by histopathological scoring of the thymus and

bursa (1 �normal; 2 �mild lymphocytic depletion; 3 �moderate

depletion; 4 � severe depletion). In addition, during all IBV passages

in both Experiment 1 and Experiment 2 tear samples obtained 16 days

after IBDV and/or CAV inoculation were diluted 1:10 and tested for

CAV and IBDV antibodies using commercially available enzyme-linked

immunosorbent assay kits as recommended by the manufacturer

(IDEXX Laboratories Inc, Westbrook, Maine, USA). The results

were expressed as S/N or S/P ratios as recommended by the

manufacturer. Induction of specific antibodies was interpreted as

successful replication of the immunosuppressive viruses. On day 15 of

age all chickens were inoculated with the ArkDPI-derived IBV vaccine

strain (100 ml containing 7.9�105 median embryo infective dose) as

follows: 25 ml/each eye and 25 ml/each nostril. Tear fluids of IBV-

infected chickens have been shown to contain high IBV levels (van

Ginkel et al., 2008). Thus, 8 days after IBV inoculation tears were

collected from all chickens in each group as described elsewhere (Toro

et al., 1993) for subsequent passaging. Further passages were performed

using tears pooled for each group, diluted in 1:2 in tryptose broth, and

ocularly inoculated (50 ml tear suspension) to the next generation of 15-

day-old immunocompetent or immunodeficient chicken groups. During

each passage, RNA was extracted from individual tear samples

(passages 1 to 5) or pooled tear samples for each group (passages 6 to

9) for RT-PCR amplification of an approximately 751-nucleotide

portion of the IBV S1 gene to confirm the presence of IBV in the tears

used for further passaging. The S1 cDNA was sequenced to confirm the

presence of IBV, to determine which vaccine subpopulations were

present in individual chickens and to identify possible additional

nucleotide changes in the S1 gene. Relative IBV genome levels in

RNA extracted from pooled tears of each group following each passage

were determined by quantitative RT-PCR (qRT-PCR) with primers and

probes designed to amplify and detect a portion of the IBV nucleo-

capsid gene as previously described (van Ginkel et al., 2008). The lower

limit of detection of this qRT-PCR assay is approximately 100 IBV

genomes/ml tears.

Experiment 2. Experiment 2 was carried out identically to Experiment

1, except that we used a different IBV vaccine lot (obtained from the

same manufacturer) and chickens were inoculated with 3.2�105 median

embryo infective dose for the first passage.

Viral RNA amplification by RT-PCR. Viral RNA was extracted from

individual tear samples obtained from each chicken of all groups using

the Qiagen QIAmp viral RNA mini kit (Qiagen, Valencia, California,

USA) following the manufacturer’s protocol. RT-PCR was performed

using the Qiagen one step RT-PCR kit (Qiagen) and previously

described primers S17F and S18R (Gallardo et al., 2010). Amplicons

were visualized by gel green staining (Phoenix Research, Candler, North

Carolina, USA) after agarose gel electrophoresis.

S1 sequencing and subpopulation identification. Amplified cDNA was

purified using the QIAquick PCR purification kit (Qiagen) and

submitted to the Massachusetts General Hospital DNA core facility

for sequencing using S1R primer (van Santen & Toro, 2008). Sequences

were aligned using Mac Vector 10.6.0 software (Mac Vector Inc., Cary,

North Carolina). Sequences were used to determine the presence of the

major vaccine population prior to inoculation into chickens, vaccine

subpopulations C1 to C5 previously found to be selected in chickens

(Gallardo et al., 2010), or novel populations. Sequence chromatograms

were analysed for the presence of subpopulation diversity. When

sequence heterogeneity was present, the simplest mixture of subpopula-

tions that would explain the chromatogram was deduced, based on the

known sequences of previously identified subpopulations of this vaccine

selected in individual chickens (Gallardo et al., 2010). For example, a

chromatogram showing both A and G at nucleotide position 167 and

both A and G at nucleotide position 388 would be interpreted to

indicate a mixture of populations C1 and C4 (see Table 1). For

sequences showing readily detectable levels of more than one popula-

tion, each subpopulation detected was recorded as present. The

incidence of each subpopulation in each group of chickens (the number

of chickens in which the subpopulation was detected divided by the

number of chickens) at each passage was determined. Sequences of the

entire S gene of IBV in pooled tears obtained after the two highest

passages in CAV-infected chickens in Experiment 1 and in IBDV-infected

chickens in Experiment 2 were obtained by sequencing RT-PCR

452 R. A. Gallardo et al.

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13

products generated using the appropriate previously described primers

(van Santen & Toro, 2008).

Results

CAV and IBDV replication. During the first IBV passage,the effects of CAV and IBDV replication were confirmedin five chickens from each group 10 days after CAV orIBDV inoculation. CAV-inoculated chickens showedreduced haematocrit values with a mean of 26 (valuesvarying between 17 and 32) compared with the meanof 33 in control birds. At necropsy, birds showed thymusatrophy. Histopathological analyses revealed severe lym-phocyte depletion in the thymic cortex with an averagescore of 4. IBDV-inoculated birds showed diffuselymphocytic depletion in the bursa consistent with thepost-necrotizing phase of IBD with mild follicularrestitution. In addition, pooled tear samples obtained16 days after IBDV and/or CAV inoculation during eachIBV passage showed CAV and IBDV antibodies in bothExperiments 1 and 2 (Figure 1). The induction ofspecific antibodies demonstrated that both CAV andIBDV replicated as expected in the inoculated chickensduring all IBV passages.

IBV RNA detection by RT-PCR during serial passages inimmunocompetent or immunodeficient chickens. In Ex-periment 1, the S1 gene of IBV was consistently amplifiedfrom tear fluid in every passage up to passage 5 in allCAV-infected immunodeficient chickens and continuedto be readily detectable in pooled tears of CAV-infectedchickens until passage 9. In contrast, IBV S1 detection inthe immunocompetent, IBDV and CAV�IBDV chickengroups had declined to 10% after passage 4 and 20% and10% after passage 3, respectively. IBV RNA was unde-tectable in these groups by passages 4 or 5 (Figure 2a).

Although IBV RNA was readily detected by conven-tional RT-PCR in tears from 90 to 100% of chickens ineach group following the first IBV passage and in pooledtears from each group used as inoculum for the secondpassage, IBV RNA could be detected and quantitated inExperiment 1 by our less sensitive qRT-PCR assay inpooled tears of only the CAV�IBDV group. IBVgenomes remained below detection levels in the immu-nocompetent and IBDV groups throughout subsequentpassages, and the levels of IBV RNA in pooled tears ofthe CAV�IBDV group decreased approximately seven-

fold between the first and second passages and fell belowthe detection limit of the qRT-PCR assay by the thirdpassage. These results are consistent with the decliningincidence of IBV-positive birds in those groups. Incontrast, in pooled tears of CAV-infected chickens,IBV levels rose over 10-fold to become quantifiable byqRT-PCR following the second passage, and increased afurther 10-fold following the third passage. However,following the fifth passage, IBV genomes fell below thedetection limit of the qRT-PCR assay, and continued tobe below the detection limit. (The qRT-PCR data for theCAV group following the fourth passage are not avail-able for technical reasons.)

Somewhat different results were obtained in Experi-ment 2. As in Experiment 1, IBV persisted in one of theimmunodeficient groups until passage 9, while beingeliminated from other groups by the fourth or fifthpassages. A difference was that IBV was maintained inthe IBDV-infected chickens rather than the CAV-infectedchickens. IBV S1 was detected by RT-PCR in the IBDVgroup up to passage 5, with detection rates varying from80 to 100%, and was readily detectable in pooled tears ofthis group until passage 9. IBV genomes in pooled tearsof the IBDV group following the second passage were15-fold higher than following the first passage andremained relatively high (levels similar to the thirdpassage in the CAV group in Experiment 1) followingthe third passage. In subsequent passages, IBV genomelevels decreased and were below the detection limit of theqRT-PCR assay following the seventh passage. On theother hand, immunocompetent, CAV and CAV�IBDVchicken groups cleared IBV by the fourth or fifthpassages, with IBV S1 detection rates during theprevious passage (before clearance) of 20%, 20% and10% for each group, respectively (Figure 2b). IBVgenome levels in pooled tears fell below the detectionlimit of the qRT-PCR assay following the second passagein the immunocompetent and CAV-inoculated groupsand following the third passage in the CAV�IBDVgroup, consistent with the declining incidence of IBV-positive birds in those groups.

IBV S1 gene sequences in chickens. IBV-Ark found intear fluids of individual chickens exhibited differences inS1 gene consensus sequences. Considering only differencesrepresenting non-synonymous changes, five distinctvirus predominant populations, all differing from the

Table 1. Differences in amino acids encoded by the S1 gene of Ark-type IBV vaccine and predominant populations detected in tears of

chickens.

Nucleotide position 127 167 226 355 388 511 593 637

Designation Amino acid position 43 56 76 119 130 171 198 213

Vaccine Nucleotide T A C T A T A T

Amino acid Tyr Asn Leu Ser Ser Tyr Lys Ser

C1 Nucleotide C A C T G T A G

Amino acid His Asn Leu Ser Gly Tyr Lys Ala

C2 Nucleotide C A C T A T A G

Amino acid His Asn Leu Ser Ser Tyr Lys Ala

C3 Nucleotide C A T C A C C G

Amino acid His Asn Phe Pro Ser His Thr Ala

C4 Nucleotide C G C T A T A G

Amino acid His Ser Leu Ser Ser Tyr Lys Ala

C5 Nucleotide T A C T A T A G

Amino acid Tyr Asn Leu Ser Ser Tyr Lys Ala

Effects of immunodeficiency on IBV drift 453

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13

major vaccine population prior to inoculation, were

found in both immunocompetent and immunodeficient

birds (Table 1). The emergent IBV subpopulations are

identical to those previously characterized and desig-

nated C1, C2, C3, C4, and C5 (Gallardo et al., 2010).

Predicted phenotypic differences between subpopula-

tions encompassed both quantitative (from one to up

to six amino acid changes) and qualitative (biochemi-

cally distant) amino acid substitutions in the portion of

S1 shown in Table 1. Corroborating previous findings,

these subpopulations were positively selected during the

first passage in chickens, while the vaccine-predominant

population was rapidly and negatively selected. Interest-

ingly, negative selection occurred both in immunocom-

petent and immunodeficient chickens. The presence of

viral populations with identical S1 sequences in multiple

chickens in this study, as well as in our previous study,

indicates that they probably emerged by selection ofvirus subpopulations already existing in the vaccinerather than arising by mutation. No additional popula-tions that had not previously been identified in chickensvaccinated with this vaccine became predominant in anyindividual chickens or groups over the course of thestudy. Of a total of 166 S1 sequences representing IBVpresent in individual chickens, 90% each indicated asingle predominant S1 sequence, without detectableamounts of different S1 sequences. An additional 2%of sequences indicated mixed populations where compo-nents of the mixture differed at only one nucleotideposition in the S1 sequence, while the remaining 8%indicated mixed populations where components of themixture differed at two or more positions. The S1sequences of the IBV present in tears of 92% of thechickens could thus be determined unequivocally.

Incidence of subpopulations by passage. Incidences afterpassage 1. Characterization of the S1 gene of IBV intears of individual chickens following the first IBVpassage of Experiments 1 and 2 consistently showedthe emergence of the distinct subpopulations describedabove (Figure 3 and Figure 4). For example, inimmunocompetent chickens (Experiment 1) subpopula-tion C1 was most frequently selected (67%) but other

S/N

rat

io

CtrCAV

CAV+IBDV Ctr

CAV

CAV+IBDV

0.0

0.5

1.0

1.5 Experiment 1 Experiment 2

CAV

* * * *

(a)

S/P

rat

io

CtrIB

DV

CAV+IBDV Ctr

IBDV

CAV+IBDV

0.0

0.5

1.0

1.5

2.0 Experiment 1 Experiment 2IBDV

* * * *

(b)

Figure 1. Detection of CAV-specific and IBDV-specific anti-

bodies in lachrymal fluid of chickens during serial IBV passages.

Antibody levels in tears collected from all groups of chickens 16

days after CAV and/or IBDV inoculation and diluted 1:10 were

determined by enzyme-linked immunosorbent assay (ELISA;

Idexx, Westbrook, Maine, USA). 1a: A competitive ELISA kit

was used for CAV antibodies; lower S/N values indicate higher

antibody levels. 1b: A direct ELISA kit was used for IBDV

antibodies; higher S/P values correspond to higher antibody

levels. Boxes: 25th percentile, median, 75th percentile. Whiskers:

minimum and maximum. *Significant differences (P B0.0001)

compared with uninoculated control birds.

0 1 2 3 4 5 60

20

40

60

80

100

ImmunocompetentIBDVCAVCAV+IBDV

Passage in chickens

Per

cent

chi

cken

s IB

V p

ositi

ve(a)

0 1 2 3 4 5 60

20

40

60

80

100

ImmunocompetentIBDVCAVCAV+IBDV

Passage in chickensP

erce

nt c

hick

ens

IBV

pos

itive

(b)

Figure 2. Percentage of IBV-positive chickens after serial

passages in immunocompetent or chickens previously inoculated

(n �10/group) at 7 days of age with CAV and/or IBDV. All

chickens were inoculated with the ArkDPI-derived IBV vaccine

strain on day 15 of age. Eight days after IBV inoculation, tears

were collected from all chickens of each group for S1 RT-PCR

detection, confirmed by sequencing, and subsequent ocular

passaging. 2a: Immunocompetent, CAV, IBDV, and

CAV� IBDV chicken groups in Experiment 1. 2b: Immunocom-

petent, CAV, IBDV, and CAV� IBDV chicken groups in

Experiment 2.

454 R. A. Gallardo et al.

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13

predominant populations included C4 (33%) and C5(17%). In another example, B-cell-deficient chickens(IBDV-infected) after passage 1 showed C2, C4, andC1 in 30% of the birds and C5 in 20%, with no birdsshowing the vaccine major population. In contrast toExperiment 1, in immunocompetent chickens of Experi-ment 2 the most frequently selected subpopulation wasC4 (60%), followed by C1 (30%). Although the incidenceof each subpopulation after passage 1 differed betweenExperiments 1 and 2, C1 and C4 showed high rates ofselection in both experiments in immunocompetentbirds, suggesting increased fitness of these two subpo-pulations (Figure 3 and Figure 4). In Experiment 2, B-cell-deficient chickens showed C4 (80%), C2 (20%), andC1 (10%). Similarly to the IBDV group of Experiment 1(shown above), C4 and C2 were among the mostfrequently selected populations in this environment.The rank of each subpopulation by incidence in eachexperiment is shown in Table 2. From the data shown inthis table it becomes clear that C4 seems to be the mostsuccessful population in all environments tested herein;that is, C4 was the most frequently selected populationin all groups in Experiment 2 and in all except theimmunocompetent group in Experiment 1, where it wasthe second most frequently selected population.

Incidence after further passages. A distinct trend wasobserved after the second and further passages in allgroups of both experiments. IBV was cleared fromimmunocompetent chickens during the fifth passage,regardless of the selected viral population predominatingin the birds. Similarly, both experiments showed thatIBV was no longer detectable after four to five passagesin chickens inoculated with IBDV�CAV. Only in thosegroups in which C2 was selected did IBV persistsuccessfully throughout the whole experimental periodof nine passages. Notably, in both cases where subpo-pulation 2 was established and persisted, it was not themost prevalent during the first passage. Interestingly,persisting C2 was selected in CAV-infected chickens inExperiment 1 and in IBDV-infected chickens in Experi-ment 2. The sequence of the entire S gene of the C2vaccine subpopulation established in immunodeficientchickens confirmed that the S1 portion of the subpopu-lations established in both experiments were identical;the subpopulations differed in only one amino acidposition in the S2 portion. In immunocompetent chick-ens of both experiments, C1 became predominant andwas cleared after the fourth passage.

Differences between the results of the two experimentsinclude the fact that one predominant population was

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hick

ens

with

sub

popu

latio

n

CAV

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hick

ens

with

sub

popu

latio

n

Immunocompetent

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hick

ens

with

sub

popu

latio

n

IBDV

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hick

ens

with

sub

popu

latio

n

IBDV+CAV

Figure 3. Incidence of IBV subpopulations in immunocompetent, IBDV-infected, CAV-infected, and CAV� IBDV-infected chicken

groups after serial passages in Experiment 1. During each passage, RNA was extracted from the individually collected tear samples for

RT-PCR amplification of a portion of the IBV S1 gene to confirm the presence of IBV in the tears. The S1 cDNA obtained was sequenced

to identify nucleotide changes in the S1 gene, and sequence chromatograms were analysed for the presence of subpopulations C1 to C5

(Gallardo et al., 2010) and the major vaccine population prior to passage in chickens (Vaccine). Some sequences, while clearly indicating

that IBV was present, were not of sufficient quality in portions of the sequence to allow unequivocal subpopulation assignment and are not

included.

Effects of immunodeficiency on IBV drift 455

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13

ultimately established in all groups in Experiment 1. Incontrast, in Experiment 2 a clearly predominant popula-tion (C2) was only selected in IBDV-infected chickens. Inall other groups in which C1 was more frequent, IBVwas cleared from the chickens between the fourth andfifth passages.

Discussion

Different portions of the IBV genome and the pheno-type contribute to the evolutionary success of this virus.For example, the replicase proteins in open readingframe 1ab have been shown to be associated withpathogenicity (Ammayappan et al., 2009). However,the spike gene in IBV appears to be the most reliablemeasure of genetic change leading to the emergence of

new viruses capable of causing disease (McKinley et al.,2011). Indeed, our previous results have confirmed thatpopulation analyses based on S1 genotype/phenotypeallow assessment of IBV drift (Gallardo et al., 2010).

We used an experimental model to examine the effectsof viral immunodeficiency on IBV evolution. In thismodel, IBV contained in tears was serially passaged inimmunocompetent or immunodeficient chickens. Weused tears because this fluid contains a high concentra-tion of IBV and it is probably a very important source forrespiratory virus spreading naturally. Indeed, lachrymalfluid drains directly into the nostrils. Thus, sneezing aswell as the cleaning habits of birds (rubbing heads on theplumage) provide optimal circumstances for respiratoryviruses contained in tears to disseminate among birds.It was fascinating to observe that, based on S1 genotype/phenotype, exactly the same IBV subpopulations

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hic

ken

s w

ith

su

bp

op

ula

tio

n Immunocompetent

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hic

ken

s w

ith

su

bpo

pu

lati

on

IBDV

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hic

ken

s w

ith

su

bp

op

ula

tio

n CAV

0 1 2 3 4 5 60

20

40

60

80

100

C1C2C3C4C5VACCINE

Passage in chickens

% c

hic

ken

s w

ith

su

bp

op

ula

tio

n

CAV+IBDV

Figure 4. Incidence of IBV subpopulations in immunocompetent, IBDV-infected, CAV-infected, and CAV� IBDV-infected chicken

groups after serial passages in Experiment 2. During each passage, RNA was extracted from the individually collected tear samples for

RT-PCR amplification of a portion of the IBV S1 gene to confirm the presence of IBV in the tears. The S1 cDNA obtained was sequenced

to identify nucleotide changes in the S1 gene, and sequence chromatograms were analysed for the presence of subpopulations C1 to C5

(Gallardo et al., 2010) and the major vaccine population prior to passage in chickens (Vaccine). Some sequences, while clearly indicating

that IBV was present, were not of sufficient quality in portions of the sequence to allow unequivocal subpopulation assignment and are not

included.

Table 2. Ranking of each subpopulation based on incidence (high to low) in immunocompetent and immunodeficient (IBDV infected,

CAV infected, CAV� IBDV infected) chickens in the first passage of Experiments 1 and 2.

Group Experiment 1 Experiment 2

Immunocompetent C1a�C4 �C5 C4 �C1

IBDV C1 �C4 �C2 �C5 C4 �C2 �C1

CAV C1 �C4 �C2 �C5 C4 �C1 �C2

CAV� IBDV C4 �C5 �C1 �Vxb C4 �C1 �C5 �C3 �C2

aSelected subpopulations, designated C1 to C5. bVaccine major population.

456 R. A. Gallardo et al.

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13

previously detected in immunocompetent chickens(Gallardo et al., 2010) emerged both in immunocompe-tent and immunodeficient chickens during the currentexperiments. This result suggests that the generation ofgenetic diversity in IBV is constrained and that selectionplays an important role. This assumption is enforced bythe fact that the same populations arose even whendifferent ArkDPI vaccine lots (although from the samemanufacturer) were used in Experiments 1 and 2. Thisfinding constitutes further evidence for phenotypic driftoccurring mainly as a result of selection. Even after ninepassages, no new mutations were selected in the S1 gene,the most variable gene of IBV. However, while we did notobserve changes as a result of point mutations, recombi-nation cannot be ruled out. For example, the C2 S1sequence might have been generated by recombinationbetween C1 and C4 S1 sequences during the first passage.

Another interesting result involves the emergence of avariety of subpopulations during the first passage, but aquick establishment of one predominant populationafter further passages. This phenomenon can be ex-plained by the fact that more replication cycles shouldresult in the selection of the populations showingincreased adaptedness.

The genetic diversity in viral isolates of immunodefi-cient individuals is attributed to shifts in populationequilibrium of the replicating viral genomes in theabsence of immune selection pressure (Rocha et al.,1991). CAV targets lymphoblasts of the thymic cortexin young chickens, causing T-cell immunodeficiency(Adair et al., 1991; Adair, 2000). T-cell responses(cytotoxic T lymphocytes) are relevant in the clearanceof IBV (Seo & Collisson, 1997; Collisson et al., 2000).CAV also reduces mucosal immune responses to IBV(van Ginkel et al., 2008). On the other hand, IBDVaffects the bursa of Fabricius, causing B-cell immuno-deficiency (Rosenberger & Gelb, 1978; Cloud et al.,1992), and antibodies also participate in the immuneresponse against IBV (for example, Koch & Kant, 1990).We previously showed that Ark-type IBV persists forlonger periods in chickens suffering from viral immuno-deficiency (Toro et al., 2006). In addition to the well-known effects of CAV and IBDV infection on acquiredimmunity, CAV and IBDV infection also have effects oninnate immunity relevant to IBV infection. IBV infectionactivates expression in chickens of numerous mRNAs asso-ciated with innate immunity, including IL-1b mRNA andmRNAs associated with type I interferon expression (Guoet al., 2008). CAV infection reduces IL-1b production bymacrophages (McConnell et al., 1993a, b). Chickensinfected with either CAV or IBDV have reduced inter-feron-alpha and interferon-gamma mRNA transcriptionin blood cells stimulated with inactivated Newcastledisease virus (Ragland et al., 2002).

Specific (acquired) immune activity against IBVdevelops very early after infection in immunocompetentchickens. IBV-specific cytotoxic T lymphocytes aredetectable as early as 3 days post inoculation (Seo &Collisson, 1997). IBV-specific IgA is detectable in tearsas early as 4 days post immunization via the ocular route(Toro et al., 1996, 1997). We previously demonstratedthat while IBV-specific IgA was detectable in lachrymalfluid of immunocompetent chickens at 8 days postinoculation it was either not detectable or present atbarely detectable levels in chickens infected with IBDVand/or CAV (van Santen et al., 2006). IBV collected

from immunocompetent chickens at 8 days post inocu-lation would thus have been subjected to selectivepressure from both innate and acquired immunity andthese pressures would have differed in chickens infectedwith IBDV and/or CAV. According to Kilbourne (1994,p. 262): ‘‘if the restricting effects of the host immuneresponse are removed, the virus has the opportunity ofemergence and, testing of mutants previously absent orsuppressed, antigenic changes can be seen related toaltered replication properties of the virus’’. The currentresults showed that only C2 successfully became estab-lished and only in either CAV-infected or IBDV-infectedchickens. These results indicate that selection indeeddoes not cease in immunodeficient chickens. The factthat C2 but no other subpopulation became establishedin immunodeficient hosts suggests a distinct adaptationof this phenotype to this type of environment. We wouldspeculate that under such circumstances other selectiveforces, such as, for example, the affinity of the S proteinto the cell receptors, may augment their selectivepreponderance. It is also certainly possible that differ-ences in regions of the genome other than the S gene thataffect replication and defence from innate and adaptiveimmune responses result in differential selection ofsubpopulations in immunodeficient and immunocompe-tent chickens.

A considerable percentage of commercial chickenssuffer from CAV and/or IBDV immunodeficiency(Hoerr, 2010). Concurrent infection with IBV is common(Toro et al., 2006). From an applied perspective, ourcurrent results indicate that such circumstances maycontribute to the emergence of novel IBV strains.

Immunocompetent chickens successfully cleared IBVafter only a few serial passages. Subpopulations C1 orC4, which became established in these birds, were not fitenough to withstand this environment and becameextinct. These birds were maintained in a controlledenvironment and the viruses did not have opportunityfor recombination with other IBV strains. Thus, genera-tion of diversity by mutation is not as frequent asanticipated (discussed above) and IBV becomes extinctunder these conditions.

The CAV�IBDV-infected chicken group producedintriguing results; that is, in neither experiment did anIBV population become established. It is possible thatthis model is inadequate to understand the drift of IBVbecause the incubation periods of CAV and IBDV differ.Indeed, IBDV is expected to produce the most severechanges in the bursa around 3 to 5 days after inoculation(Pope, 1996), while CAV induces maximal changes in thethymus a few days later (around 10 days after parenteralinoculation; van Santen et al., 2004). Co-infection withCAV and IBDV may thus lead to different timing andkinetics of innate immune responses than infection witheither virus alone, providing an altered environment tothe replicating IBV strain.

Acknowledgements

The authors thank Cassandra Breedlove, Lisa Parsons,Stephen Kitchens, Natalia Petrenko, and Stephen Gulleyfor technical assistance. Support for this work wasprovided by United States Department of AgricultureCooperative State Research, Education, and ExtensionService NRI # 2007-35204-18330.

Effects of immunodeficiency on IBV drift 457

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13

References

Adair, B.M. (2000). Immunopathogenesis of chicken anaemia virus

infection. Developmental and Comparative Immunology, 24, 247�255.

Adair, B.M., McNeilly, F., McConnell, C.D., Todd, D., Nelson, R.T. &

McNulty, M.S. (1991). Effects of chicken anaemia agent on

lymphokine production and lymphocyte transformation in experi-

mentally infected chickens. Avian Diseases, 35, 783�792.

Ammayappan, A., Upadhyay, C., Gelb, J. & Vakharia, V.N. (2009).

Identification of sequence changes responsible for the attenuation of

avian infectious bronchitis virus strain Arkansas DPI. Archives of

Virology, 154, 495�499.

Cavanagh, D. (1981). Structural polypeptides of coronavirus IBV.

Journal of General Virology, 53, 93�103.

Cavanagh, D. (1983). Coronavirus IBV: structural characterization of

the spike protein. Journal of General Virology, 64, 2577�2583.

Cavanagh, D. (1984). Structural characterization of IBV glycoproteins.

Advances in Experimental Medicine and Biology, 173, 95�108.

Cavanagh, D. & Davis, P.J. (1986). Coronavirus IBV: removal of spike

glycopolypeptide S1 by urea abolishes infectivity and haemagglutina-

tion but not attachment to cells. Journal of General Virology, 67,

1443�1448.

Cloud, S.S., Lillehoj, H.S. & Rosenberger, J.K. (1992). Immune

dysfunction following infection with chicken anaemia agent and

infectious bursal disease virus. I. Kinetic alterations of avian

lymphocyte subpopulations. Veterinary Immunology and

Immunopathology, 34, 337�352.

Collisson, E.W., Pei, J., Dzielawa, J. & Seo, S.H. (2000). Cytotoxic T

lymphocytes are critical in the control of infectious bronchitis virus in

poultry. Developmental and Comparative Immunology, 24, 187�200.

Gallardo, R.A., van Santen, V.L. & Toro, H.E. (2010). Host intraspatial

selection of infectious bronchitis virus populations. Avian Diseases,

54, 807�813.

Ginkel, F.W. van, Santen, V.L. van, Gulley, S.L. & Toro, H. (2008).

Infectious bronchitis virus in the chicken Harderian gland and

lachrymal fluid: viral load, infectivity, immune cell responses, and

effects of viral immunodeficiency. Avian Diseases, 52, 608�617.

Guo, X., Rosa, A.J.M., Chen, D.-G. & Wang, X. (2008). Molecular

mechanisms of primary and secondary mucosal immunity using

avian infectious bronchitis virus as a model system. Veterinary

Immunology and Immunopathology, 121, 332�343.

Hoerr, F.J. (2010). Clinical aspects of immunosuppression in poultry.

Avian Diseases, 54, 2�15.

Kilbourne, E.D. (1994). Host determination of viral evolution: a

variable tautology. In S.S. Morse (Ed.). The Evolutionary Biology of

Viruses (pp. 253�271). New York: Raven.

Koch, G., Hartog, L., Kant, A. & Van Roozelaar, D.J. (1990). Antigenic

domains on the peplomer protein of avian infectious bronchitis virus:

correlation with biological functions. Journal of General Virology, 71,

1929�1935.

Koch, G. & Kant, A. (1990). Binding of antibodies that strongly

neutralise infectious bronchitis virus is dependent on the glycosyla-

tion of the viral peplomer protein. Advances in Experimental

Medicine and Biology, 276, 143�150.

Kusters, J.G., Jager, E.J., Niesters, H.G.M. & Van der Zeijst, B.A.M.

(1990). Sequence evidence for RNA recombination in field isolates of

avian coronavirus infectious bronchitis virus. Vaccine, 8, 605�608.

Kusters, J.G., Niesters, H.G.M., Bleumink-Pluym, N., Davelaar, F.G.,

Horzinek, M.C. & Van der Zeijst, B.A.M. (1987). Molecular

epidemiology of infectious bronchitis virus in the Netherlands.

Journal of General Virology, 68, 343�352.

Kusters, J.G., Niesters, H.G.M., Lenstra, J.A., Horzinek, M.C. & Van

der Zeijst, B.A.M. (1989). Phylogeny of antigenic variants of avian

coronavirus IBV. Virology, 169, 217�221.

McConnell, C.D., Adair, B.M. & McNulty, M.S. (1993a). Effects of

chicken anaemia virus on cell-mediated immune function in chickens

exposed to the virus by a natural route. Avian Diseases, 37, 366�374.

McConnell, C.D., Adair, B.M. & McNulty, M.S. (1993b). Effects of

chicken anaemia virus on macrophage function in chickens. Avian

Diseases, 37, 358�365.

McKinley, E.T., Hilt, D.A. & Jackwood, M.W. (2008). Avian corona-

virus infectious bronchitis attenuated live vaccines undergo selection

of subpopulations and mutations following vaccination. Vaccine, 26,

1274�1284.

McKinley, E.T., Jackwood, M.W., Hilt, D.A., Kissinger, J.C.,

Robertson, J.S., Lemke, C. & Paterson, A.H. (2011). Attenuated live

vaccine usage affects accurate measures of virus diversity and

mutation rates in avian coronavirus infectious bronchitis virus.

Virus Research, 158, 225�234.

McNulty, M.S. & Todd, D. (2008). Chicken anaemia virus. In

L. Dufour-Zavala (Ed.). Isolation, Identification, and Characterization

of Avian Pathogens, 5th edn. (pp. 124�127). Athens, GA: The

American Association of Avian Pathologists.

Mockett, A.P.A., Cavanagh, D. & Brown, T.D.K. (1984). Monoclonal

antibodies to the S1 spike and membrane proteins of avian infectious

bronchitis coronavirus strain Massachusetts M41. Journal of General

Virology, 65, 2281�2286.

Pope, C.R. (1996). Lymphoid system. In C. Riddell (Ed.). Avian

Histopathology (pp. 17�44). Kennett Square, PA: New Bolton Center,

AAAP.

Ragland, W.L., Novak, R., El-Attrache, J., Savic, V. & Ester, K. (2002).

Chicken anaemia virus and infectious bursal disease virus interfere

with transcription of chicken IFN-alpha and IFN-gamma mRNA.

Journal of Interferon and Cytokine Research, 22, 437�441.

Rocha, E., Cox, N.J., Black, R.A., Harmon, M.W., Harrison, C.J. &

Kendal, A.P. (1991). Antigenic and genetic variation in influenza a

(H1N1) virus isolates recovered from a persistently infected immu-

nodeficient child. Journal of Virology, 65, 2340�2350.

Rosenberger, J.K. & Gelb, J. Jr. (1978). Response to several avian

respiratory viruses as affected by infectious bursal disease virus. Avian

Diseases, 22, 95�105.

Santen, V.L. van, Joiner, K.S., Murray, C., Petrenko, N., Hoerr, F.J. &

Toro, H. (2004). Pathogenesis of chicken anaemia virus: comparison

of the oral and the intramuscular routes of infection. Avian Diseases,

48, 494�504.

Santen, V.L. van & Toro, H. (2008). Rapid selection in chickens of

subpopulations within ArkDPI-derived infectious bronchitis virus

vaccines. Avian Pathology, 37, 293�306.

Santen, V.L. van, Toro, H., Ginkel van, F.W. Joiner, K.S. & Hoerr, F.J.

(2006). Effects of chicken anaemia virus and/or infectious bursal

disease virus on infectious bronchitis virus infection and immune

responses. In U. Heffels-Redmann & E.F. Kaleta (Eds.). V. Interna-

tional Symposium on Avian Corona- and Pneumoviruses and Compli-

cating Pathogens (pp. 296�299). Rauischholzhausen, Germany.

Seo, H.S. & Collisson, E.W. (1997). Specific cytotoxic T lymphocytes are

involved in in vivo clearance of infectious bronchitis virus. Journal of

Virology, 71, 5173�5177.

Toro, H., Espinoza, C., Ponce, V., Rojas, V., Morales, M.A. & Kaleta, E.F.

(1997). Infectious bronchitis: effect of viral doses and routes on

specific lacrimal and serum antibody responses in chickens. Avian

Diseases, 41, 379�387.

Toro, H., Lavaud, P., Vallejos, P. & Ferreira, A. (1993). Transfer of IgG

from serum to lachrymal fluid in chickens. Avian Diseases, 37, 60�66.

Toro, H., Reyes, E., Redmann, T. & Kaleta, E.F. (1996). Local and

systemic specific antibody response of different chicken lines after

ocular vaccination against infectious bronchitis. Journal of Veterinary

Medicine B, 43, 449�454.

Toro, H., van Santen, V.L., Hoerr, F.J. & Breedlove, C. (2009). Effects of

chicken anaemia virus and infectious bursal disease virus in

commercial chickens. Avian Diseases, 53, 94�102.

Toro, H., van Santen, V.L., Li, L., Lockaby, S.B., van Santen, E. &

Hoerr, F.J. (2006). Epidemiological and experimental evidence for

immunodeficiency affecting avian infectious bronchitis. Avian

Pathology, 35, 455�464.

458 R. A. Gallardo et al.

Dow

nloa

ded

by [

134.

117.

10.2

00]

at 0

6:00

30

Sept

embe

r 20

13