RESEARCH ARTICLE

Phylogenetic Analysis of Anopheles (Cellia) subpictus Grassi UsingrDNA-ITS2 Sequence

Jainder S. Chhilar • Sudarshan Chaudhry

Received: 6 February 2011 / Revised: 22 March 2012 / Accepted: 27 March 2012 / Published online: 11 April 2012

� Zoological Society, Kolkata, India 2012

Abstract Anopheles subpictus Grassi is one of the most

abundant malaria vector mosquitos in Indian subcontinent

especially in post monsoon months. This taxon has been

speculated to be composed of four sibling species recog-

nised using morphological and cytogenetic parameters

provisionally named A, B, C and D. One of the sibling

species ‘B’ has well documented status as a vector of

malaria parasite in the Oriental region. Molecular phylo-

genetic analysis of this mosquito was done to discern sib-

ling/cryptic species using internal transcribed spacer 2

region of the nuclear ribosomal DNA (rDNA-ITS2). An.

culicifacies sibling species A, a member of Myzomia ser-

ies, was used as an out-group to root the trees. In the

present analysis rDNA-ITS2 region was PCR amplified,

sequenced and the sequences obtained were then subjected

to phylogenetic analysis with the sequences already present

in the sequence database GenBank. Multiple sequence

alignment was performed using ClustalX and further

manually annotated in MEGA 4. Initial analysis suggested

extreme 30 end sequence divergence within different pop-

ulations. Phylogenetic analysis of this spacer was done

using maximum parsimony, maximum likelihood, and

distance matrix—neighbour joining methods performed

with bootstrapped dataset that were executed in DNA-

PARS, DNAML, DNADIST and neighbour programs in

Phylip software package respectively. Phylogenetic trees

produced were similar in topology but differed in bootstrap

support. Results support the division of this taxon into at

least two major clades differing in their sequence compo-

sition and product size that are recommended to be

renamed as An. subpictus inland form and An. subpictus

coastal form. Differences include insertions/deletions,

transitions and transversions; especially one large 30 end

deletion event. The sibling species status of An. subpictus

has been analysed critically.

Keywords Anopheles subpictus � rDNA-ITS2 �Phylogeny � Sibling species

Introduction

Mosquitoes belonging to the family Culicidae, subfamily

Anophelinae act as vectors of various pathogens including

malarial parasite. For more than a century, mosquito tax-

onomists and systematists have concentrated their efforts on

the biology, identification and classification of members of

the family Culicidae attempting to determine their diversity,

vectorial status and phylogeny (Coluzzi 1970; Kitzmiller

1976; Harbach 1994; Harbach and Kitching 1998, 2005;

Sallum et al. 2002). Presently, there are 420 morphologically

distinguishable anopheline species in the world of which 70

are considered to be vectors of malaria. Out of these 420, 56

are prevalent in Indian subcontinent including 13 malaria

vectors (Knight and Stone 1977; Rao 1984; Nagpal and

Sharma 1995). Anopheles (Cellia) subpictus Grassi 1899 is

the most abundant anopheline in most parts of the Indian

subcontinent and South-east Asia (Rao 1984; Chandra et al.

2010). Its role as a vector of malaria, filariasis, and West Nile

virus has been well documented (Panicker et al. 1981; Ku-

lkarni 1983; Banerjee et al. 1991; Amerasinghe et al. 1992;

Abhyawardana et al. 1996; Sahu 1998; Amerasinghe and

J. S. Chhilar (&)

Department of Zoology, Government PG College, Gohana,

Sonipat 131301, Haryana, India

e-mail: jainder@gmail.com

S. Chaudhry

Mosquito Cytogenetic Unit, Department of Zoology, Panjab

University, Chandigarh 160014, India

123

Proc Zool Soc (Jan-June 2012) 65(1):1–10

DOI 10.1007/s12595-012-0021-8

TH

EZ

O

OLOGICAL SOC

IET

YKO LK ATA

Amerasinghe 1999; Hubalek and Halouzka 1999; WHO

1999; Chatterjee and Chandra 2000; Chandra et al. 2010).

Presently An. subpictus is a member of the Pyretophorus

series of subgenus Cellia having a sister group relationship

with An. indefinitus (Anthony et al. 1999).

To solve the problem sibling species have been created

especially in vector species discrimination, various tools

and techniques were successfully used including morphol-

ogy, polytene chromosome comparative cytogenetics, and

molecular level tools viz. enzyme electrophoresis, cuticular

hydrocarbons and DNA based markers resulting in identi-

fication of more than 30 different sibling species complexes

in The genus Anopheles (Coluzzi 1970; Green et al. 1985;

Pape 1992; Subbarao and Sharma 1997; Xu et al. 1998;

Marrelli et al. 1999; Huong et al. 2001; Linton et al. 2001;

Manonmani et al. 2001; Goswami et al. 2005; Ma et al.

2006; Foley et al. 1998, 2007). Ribosomal DNA (rDNA)

has been extensively used in the past for identification and

discrimination of sibling species in anophelines especially

the internal transcribed spacer 2 (ITS2) region (Fritz et al.

1994; Collins and Paskewitz 1996; Favia et al. 2001; Chen

et al. 2003). Also the problems and complexities in analysis

of cryptic taxa at molecular level, the systematics and

classification of anophelines, application of various

molecular phylogenetic approaches and troubleshooting

during analysis have been thoroughly reviewed (Nei 1996;

Harbach and Kitching 1998, 2005; Besansky 1999; Krzy-

winski et al. 2001; Sanderson and Bradley Shaffer 2002;

Krzywinski and Besansky 2003; Philippe et al. 2005).

In this context, based on the presence of inversion

genotypes two sibling species provisionally named A and B

were reported within An. subpictus (Suguna 1982; Reuben

and Suguna 1983). Further in 1994, Suguna and co-workers

presented morphological evidences in support of the pres-

ence of two additional species C and D raising the number of

sibling species in this taxon to four. Reports of presence of

morphological and cytogenetic polymorphism in An. sub-

pictus (Kirti and Kaur 2004; Chhilar and Chaudhry 2005)

made it necessary to have a molecular viewpoint on its

sibling species status. Recently, Chhilar (2009) and Suren-

dran et al. (2010) have reported that published morpholog-

ical characters are not enough to identify some members of

the An. subpictus complex hence the present study based on

the sequence characteristics of rDNA-ITS2 has implications

on the sibling species within the An. subpictus complex.

Materials and Methods

Collection of Mosquitoes and Identification

Wild populations of An. subpictus constituted the material

for the present study. The larvae and adults were collected

from the States of Haryana and Punjab (North-western

India) (Table 1). The larvae, in various stages of devel-

opment were segregated, put in separate enamel bowls and

fed on a protein rich diet of finely powdered dog biscuits

and yeast tablets in the ratio of 6:4 (Chaudhry et al. 2005).

For the preliminary sorting the dichotomous keys of Wattal

and Kalra (1967), Nagpal and Sharma (1995) and a rapid

field key prepared by the present authors for all the species

prevalent in the Chandigarh region (unpublished) were

followed. The final confirmation was done from the stan-

dard banding pattern of the larval salivary polytene

X-chromosome from fourth instar larvae (Chaudhry et al.

2005).

DNA Extraction and PCR Amplification

DNA was extracted from more than ten individual adult

specimens from each population by following the standard

phenol–chloroform extraction method (Sambrook et al.

1989). The wings and head of individual mosquito females

were mounted on the slides for population identification

and differentiation and the rest of the body parts were used

for DNA extraction. Primers were designed for rDNA-ITS2

spacer region using primer3 web interface software from

rDNA-ITS2 sequence (accession number AY049004)

(Table 2). These primers aligned to sites in the conserved

5.8S (ITS2FP) and 28S (ITS2RP) regions flanking the

ITS2. The working PCR reaction mixture had 19 PCR

reaction buffer containing 1.5 mM MgCl2; 0.2 mM dNTPs

and 2.5 lM of each primer. To this 1 ll of sample DNA

(diluted 50 times) and 1 ll of Taq polymerase enzyme (3

units per reaction) were also added. Thermocycler was

programmed for the PCR with the following reaction

conditions: step 1: initial denaturation at 94 �C for 4 min,

step 2: denaturation at 94 �C for 1 min, step 3: annealing at

50 �C for 1 min, step 4: amplification at 72 �C for 1 min,

step 5: repeat step 2–4 for 30 cycles, step 6: final extension

at 72 �C for 5 min. The PCR amplified products were

resolved on 2 % Agarose gels using 0.59 TBE buffer.

Table 1 Collection details of An. subpictus populations

S. no. Place of collection Longitude/latitude Date of collection Life stage of mosquito

1 Bank Colony, Rohtak Road, Bhiwani (HR) 28.46 N 76.18 E 10.10.03 Adults, larvae, pupae

2 Gohana Road at Purkhas Road diversion, Sonipat (HR) 29.00 N 70.00 E 21.09.03 Larvae

3 Kishan Pura, Patiala Road, Sangrur (PB) 30.12 N 75.53 E 2.11.03 Adults, larvae, pupae

2 Proc Zool Soc (Jan-June 2012) 65(1):1–10

123

Standard 100 bp DNA ladder was used for checking the

amplified product size. All the chemicals were purchased

from Bangalore Genei P Ltd, Bangalore, India. PCR

products were commercially sequenced from genoMbio

Technologies Pvt. Ltd., Pune, India. The PCR end product

size varied from 667 to 680 bp while the ITS2 spacer size

varied from 558 to 559 bp. Sequences were annotated

using ChromasPro version 1.41 and submitted using stand

alone sequence preparation tool ‘Sequin’ to the GenBank

database. The sequences submitted to GenBank database

were assigned accession numbers EF1601868–EF1601870.

Apart from the populations sequenced during the present

study from North-west India other sequences of rDNA-

ITS2 available at GenBank database having the acces-

sion numbers AY049004.1, AF406615.1, AF406614.1,

AF406613.1, and AF406616.2 were retrieved and used for

phylogenetic analysis (Table 3). rDNA-ITS2 sequence

having accession no AY702488.1 of An. culicifacies sibling

species A, a member of Myzomia series in subgenus Cellia,

was used as out-group to root the phylogenetic trees.

Phylogenetic Analysis

Multiple sequence alignment (MSA) was performed using

ClustalX and then it was manually edited in MEGA version

4 (Tamura et al. 2007). The MSA was obtained in a *.phy

format file to perform phylogenetic analysis with the

Phylip software package (Felsenstein 2004). For a thor-

ough Phylogenentic analysis both the distance and char-

acter based approaches—neighbour joining (NJ) analysis,

maximum-parsimony (MP) analysis, and maximum-likeli-

hood (ML) analysis respectively were preformed and

Kimura two-parameter distances were calculated using

Total substitution and transition/transversion ratio (Nei

1996; Sanderson and Bradley Shaffer 2002; Philippe et al.

2005). The bootstrapping was carried out prior to analysis

with 1000, 100, and 10000 replicates for parsimony, ML,

and NJ approach respectively. The protocols and steps

were followed from Aiyar (2000) for ClustalX and Retief

(2000) for Phylip.

Results

The MSA of annotated rDNA-ITS2 sequences of An.

subpictus populations with out-group An. culicifacies

resulted in a total of 662 alignment sites out of which the

base sequence of only 134 sites was constant (Fig. 1). In

other regions as many as 21 insertions/deletions (indels)

varying in length requiring gaps in alignment could be

identified. When compared carefully, it was found that ten

of these indels were due to sequence divergence between

the coastal populations (A, B and E) and inland populations

(F, G, H, C and D) (see Table 3 for details). Out of the ten

Table 2 rDNA-ITS2 forward and reverse primers used in the present study

Primer Sequence Properties

ITS2 FP 50-GTGAACTGCAGGACACATGAA-30 Length: 21 bases,

Tm: 59.74 �C,

GC%: 47.62

ITS2 RP 50-TGCTTAAATTTAGGGGGTAGTCA-30 Length: 23 bases,

Tm: 59.10 �C,

GC%: 39.13

Table 3 Details of ITS2 sequence composition in An. subpictus

Species/population Population code Accession no. ITS2 length GC% content

AP ITS2 AP ITS2

Anopheles subpictus/Sri Lanka coastal region A Gb/AY049004.1 618 477 53.23 55.04

Anopheles subpictus/Sri Lanka inland B Gb/AF406615.1 662 474 54.68 56.54

Anopheles subpictus/Sri Lanka inland 2 C Gb/AF406614.1 755 564 53.24 54.78

Anopheles subpictus/Sri Lanka inland 3 D Gb/AF406613.1 754 564 53.05 54.78

Anopheles subpictus/Sri Lanka coastal region 2 E Gb/AF406616.2 668 521 53.44 53.84

Anopheles subpictus/India Sonipat F Gb/EF601868 670 558 55.82 56.81

Anopheles subpictus/India Bhiwani G Gb/EF601869 682 559 55.27 56.17

Anopheles subpictus/India Sangrur H Gb/EF601870 667 558 55.47 57.52

AP amplified product, length in base pair

Proc Zool Soc (Jan-June 2012) 65(1):1–10 3

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Fig. 1 MSA of rDNA-ITS2 spacer sequences of An. subpictus using ClustalX software in Phylip (*.phy) format. Indels in alignment are denoted

by dash (–)

4 Proc Zool Soc (Jan-June 2012) 65(1):1–10

123

indels, seven were found in the 50 half of ITS2 sequences

under study. Further, one large 30 terminal indel of 121

bases between ‘541 base and 662 base’ was found in

populations A, B, and E (the coastal populations).

Phylogenetic Distance

Kimura’s two-parameter distances were calculated using

total substitutions i.e. transitions ? transversions (d =

s ? v) and transition/transversion ratio (R = s/v) using

MEGA 4 software (Table 4).

Kimura Two-Parameter Distance Using d = s ? v

As a result of the comparisons of sequences it was noticed

that the average genetic distance was 0.454 among the

members of all the populations (referred hereafter as taxa)

when all gaps were considered as complete deletions,

whereas it was 0.453 when all gaps were considered as

pair-wise deletions. The maximum and minimum distance

between different taxa was found between populations A

and E (0.000) and B and D (0.253) when all gaps were

considered as complete deletion whereas, on the basis of all

the gaps considered as pair-wise deletion, the minimum

and maximum distances were present between population

A and E (0.000) and D and E (0.301) respectively.

Kimura Two-Parameter Distance Using R (Transition/

Transversion Ratio)

In all the populations under study, the average transition/

transversion ratio was 1.246 when all gaps were considered

as complete deletions, whereas it was 1.168 when all gaps

were considered as pair-wise deletions. The minimum and

maximum ratios were observed between population G and

D (0.480) and G, A, and E (1.312) when all gaps were

considered as complete deletion, whereas with all gaps

being considered as pair-wise deletion it was between

population C and D (0.330) and G and B (1.310).

Phylogenetic Relationships

The MSA file in Phylip format called outfile was manually

edited using a standard text editor and ‘–’ or ‘.’ were

replaced by ‘?’ characters so that they are counted as a

single gap. In all the analysis no weights were used and the

gaps were excluded. The bootstrapped data was used for

analysis using DNAML, DNAPARS, and DNADIST (ML,

MP, and NJ) that resulted in consensus trees with similar

topology but varying in bootstrap support. Most of the

phylogenetically critical nodes were significantly supported

by bootstrap values of [90 % (Table 5; Figs. 2, 3, 4).

Table 4 Nucleotide P-distance matrix between rDNA-ITS2 sequen-

ces of An. subpictus populations and An. culicifacies (out-group)

using Kimura two-parameter method with d: transitions ? transver-

sions (lower-left diagonal), with R = s/v (transition/transversions)

(upper-right diagonal)

Population A B C D E F G H

A 0.000 1.182 0.897 0.884 ? 1.056 1.312 1.189

B 0.030 0.000 0.976 0.960 1.182 1.017 1.350 1.066

C 0.234 0.244 0.000 0.500 0.897 1.098 0.564 0.899

D 0.243 0.253 0.028 0.000 0.884 0.766 0.480 0.852

E 0.000 0.030 0.234 0.243 0.000 1.056 1.312 1.189

F 0.207 0.192 0.079 0.084 0.207 0.000 0.604 1.132

G 0.226 0.217 0.095 0.097 0.226 0.057 0.000 0.983

H 0.251 0.225 0.105 0.102 0.251 0.045 0.095 0.000

Table 5 Bootstrap value variation

S. no Taxon/clade MP ML NJ

1 (F ? H) 939 86 8153

2 (C ? D) 981 100 9853

3 (A ? E) 999 100 9975

4 ((F ? H) ? G) 460 – –

5 ((A ? E) ? B) 1000 86 9780

6 ((F ? H) ? (C ? D)) – 48 6830

7 ((F ? H) ? G) ? (C ? D)) 1000 – –

8 ((F ? H) ? (D ? C) ? G) – 91 9299

Fig. 2 Phylogenetic tree generated by MP An. culicifacies was used

as out-group to root the tree. Numbers on branches are bootstrap

values using 1,000 replicates. Taxa are abbreviated as in Table 3

Fig. 3 Phylogenetic tree generated by ML An. culicifacies was used

as out-group to root the tree. Numbers on branches are bootstrap

values using 100 replicates. Taxa are abbreviated as in Table 3

Proc Zool Soc (Jan-June 2012) 65(1):1–10 5

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Populations (A ? E), (D ? C), and (F ? H) clustered

together in different clades while population B clustered

with clade (A ? E) as a basal group. Population G was

found to be basal to either (F ? H) or (C ? D) clade.

Population G clustered paraphyletically in inland popula-

tions clades either as ((F ? H) ? G) in MP tree where it

was supported by a bootstrap value of 460 only, whereas it

clustered as ((F ? H) ? (D ? C) ? G) in ML and NJ

trees with significant bootstrap values of 91 and 9,299

(Table 5; Figs. 2, 3, 4).

Discussion

Variability in Length and Sequence Composition

The sequence analysis revealed two types of rDNA-ITS2

spacers in this species—one consisting of 558–564 bp

(populations F, G, H, C, and D) while the other consisting of

474–521 bp (populations A, B, and E) (Table 3). When the

length of ITS2 was reviewed within the subgenus Anoph-

eles, Maculipennis complex has an average length of

305 bp (Porter and Collins 1991) while Quadrimaculatus

complex has 305–310 bp (Cornel et al. 1996), An. petrag-

nani has 302 bp and An. claviger has 341 bp (Kampen et al.

2003). Similarly, within the subgenus Nyssorhynchus, An.

nuneztovari has a range of 363–369 bp (Fritz et al. 1994).

To the contrary the members of the subgenus Cellia have

longer ITS2 spacers for example: An. gambiae has 426 bp

(Paskewitz et al. 1993), Dirus complex has 710–716 bp,

while Punctulatus group has 549–563 bp (Beebe et al.

2000c), an exception being the Minimus group members

having ITS2 region as small as 227 bp in An. varuna and

375 bp in An. minimus C (Phuc et al. 2003). Beebe and

Cooper (2000) reported in their study on the Punctulatus

group that the 50 end sequence was more conserved than the

30 sequence, a condition which is similar to the one found in

the present populations of An. subpictus. Interestingly 30

end of the ITS2 region has been contended to be useful in

attaining a stable secondary structure (Joseph et al. 1999;

Schultz et al. 2005; Dassanayake et al. 2008).

The comparative base composition of the ITS2

sequences reveals that purines are more than pyrimidines in

all the populations (Table 6). The average base composi-

tion for An. subpictus was found to be 21.6 % of A, 22.6 %

of T, 29.5 % of G and 26.2 % of C. The MSA revealed

genomic divergence in the spacer length and base com-

position which manifested as Transitions, Transversions

and indels in the alignments. In comparison to the ampli-

fied products that included 5.8s and 28s rDNA regions

(which were annotated) the ITS2 spacer was found to be

GC rich (Tables 3, 6). When compared to other anopheline

species, the average GC content of 56.68 % is very close to

the An. gambiae complex in the same Pyretophorus series

that is having 55 % of the total GC content. However,

some of the members of subgenus Cellia have longer ITS2

spacers leading to higher GC content. For example, An.

dirus and An. punctulatus have as much as 61–71 % where

the increase is attributed to evolutionary drift and natural

selection for greater fitness and survival of the species

against the natural mutational load. There is a general

observation that, within the species complexes, the extent

of sequence variations among the members is not consis-

tent (Beebe et al. 1999, 2000a, b, c; Marinucci et al. 1999;

Beebe and Cooper 2000).

Table 6 Comparative base composition of ITS2 sequences in populations of An. subpictus

Population

code

Nucleotide frequency

of base G

Nucleotide frequency

of base C

Nucleotide frequency

of base A

Nucleotide frequency

of base T

A 30.5 24.6 20.6 24.4

B 31.4 25.1 19.6 23.8

C 28.7 26.1 22.5 22.7

D 28.0 26.8 22.3 22.9

E 29.4 24.4 21.3 24.8

F 30.3 26.5 21.7 21.5

G 29.5 26.7 21.3 22.5

H 30.5 27.1 21.0 21.5

Fig. 4 Phylogenetic tree generated by NJ with Kimura two-param-

eter distances An. culicifacies was used as out-group to root the tree.

Numbers on branches are bootstrap values using 1,000 replicates.

Taxa are abbreviated as in Table 3

6 Proc Zool Soc (Jan-June 2012) 65(1):1–10

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Phylogenetic Distance and Relationships

The analysis of Kimura two-parameter and other genetic

distances revealed that the minimum distance was

encountered between members of the inland populations of

An. subpictus and the maximum distance is between

An. subpictus coastal populations and An. culicifacies

(out-group) using both total substitution and transition/

transversion ratio approach. The same is reflected in the

phylogenetic trees derived using bootstrapped datasets

Fig. 5 ITS2 consensus

sequence of An. subpictusinland type (population C, D, F,

G, and H)

Fig. 6 ITS2 consensus

sequence of An. subpictuscoastal type (A, B, and E

populations)

Fig. 7 MSA of consensus sequences for inland type A and coastal type B (showing the large 30 indel)

Proc Zool Soc (Jan-June 2012) 65(1):1–10 7

123

(Figs. 2, 3, 4). All the three phylogenetic trees derived

from rDNA-ITS2 sequence dataset support the presence of

sibling species in An. subpictus complex as most of the

important branches have [70 % bootstrap support. When a

condensed tree for 75 % bootstrap value is considered all

the branches collapse and the populations clearly form the

two basal clades representing the inland and coastal pop-

ulations. Similarly, the inner nodes for ancestral taxonomic

units of inland populations and coastal populations are

significantly supported by bootstrap values in trees

obtained in all the phylogenetic approaches. As the prob-

ability of obtaining the correct tree topology is above 95 %

in a 500 bp length sequence using any of the MP, ML or

NJ-based approach (Nei 1996), it can be inferred that the

tree topology obtained using different approaches in the

present study is also correct.

Finally, it can be concluded that population B from Sri

Lanka is actually a member of provisionally named An.

subpictus coastal form, while the population from Bhiwani

(G) India is basal to all the inland populations representing

the inland type form. In relevance to the results discussed

in the foregoing text consensus sequences were generated

for both the inland and coastal form (for various popula-

tions as detailed in Table 3; Figs. 5, 6) of this species and

aligned using ClustalX (Fig. 7) as a ready reference stan-

dard for future studies. It is recommended that the two

sibling species A and B be renamed as inland and coastal

forms. Though, there is very little support for sibling spe-

cies C and D in the present molecular study still further

inter-laboratory joint research effort to describe proper

taxonomic status of these forms using multiple parameters

from whole of its geographical and vectorial range are

required before deciding about the status of sibling species

C and D.

Acknowledgments The authors are thankful to Chairperson,

Department of Zoology, Panjab University, Chandigarh for providing

the laboratory facilities under its special assistance programme (SAP)

and Centre of Advance Studies (CAS) of University Grants Com-

mission, New Delhi. The first author is grateful to the Council of

Scientific and Industrial Research, New Delhi for providing research

grant vide F. no. 9/135(441)/2K2/EMR-I. The authors are highly

grateful to the reviewers for their critical suggestions.

References

Abhyawardana, T.A., S.R. Wijesuriya, and R.K. Dilrukshi. 1996.

Anopheles subpictus complex: Distribution of sibling species in

Sri Lanka. Indian Journal of Malariology 33: 53–60.

Aiyar, A. 2000. The use of Clustal W and Clustal X for multiple

sequence alignment. In Methods in Molecular Biology, vol 132:Bioinformatics methods and protocols, ed. S. Misener, and S.A.

Krawetz, 221–241. Totowa, NJ: Humuna Press.

Amerasinghe, P.H., and F.P. Amerasinghe. 1999. Multiple host

feeding in field population of Anopheles culicifacies and

Anopheles subpictus in Sri Lanka. Medical and VeterinaryEntomology 13: 124–131.

Amerasinghe, P.H., F.P. Amerasinghe, R.A. Wirtz, N.G. Indrajith, W.

Somapala, L.R. Pereira, and A.M. Rathnayake. 1992. Malaria

transmission by Anopheles subpictus (Diptera: Culicidae) in a

new irrigation project in Sri Lanka. Journal of MedicalEntomology 29: 577–581.

Anthony, T.G., R.E. Harbach, and I.J. Kitching. 1999. Phylogeny of

the Pyretophorus series of Anopheles subgenus Cellia (Diptera:

Culicidae). Systematic Entomology 24: 193–205.

Banerjee, P.K., K.N. Gosh, and R.N. Chatterjee. 1991. Gel diffusion

analysis of host preference pattern of Anopheles subpictus in

West Bengal, India. Indian Journal of Malariology 28: 157–159.

Beebe, N.W., and R.D. Cooper. 2000. Systematics of malaria vectors

with particular reference to the Anopheles punctulatus group.

International Journal of Parasitology 30: 1–17.

Beebe, N.W., J.T. Ellis, R.D. Cooper, and A. Saul. 1999. DNA

sequence analysis of the ribosomal DNA ITS2 region for the

Anopheles punctulatus group of mosquitoes. Insect MolecularBiology 8: 381–390.

Beebe, N.W., R.D. Cooper, D.H. Foley, and J.T. Ellis. 2000a.

Populations of the south-west Pacific malaria vector Anophelesfarauti s.s. revealed by ribosomal DNA transcribed spacer

polymorphisms. Heredity 84: 244–253.

Beebe, N.W., R.D. Cooper, D.A. Morrison, and J.T. Ellis. 2000b.

Subset partitioning of the ribosomal DNA small subunit and its

effects on the phylogeny of the Anopheles punctulatus group.

Insect Molecular Biology 9: 515–520.

Beebe, N.W., R.D. Cooper, D.A. Morrison, and J.T. Ellis. 2000c. A

phylogenetic study of the Anopheles punctulatus group of

malaria vectors comparing rDNA sequence alignments derived

from the mitochondrial and nuclear small ribosomal subunits.

Molecular Phylogenetics and Evolution 17(3): 430–436.

Besansky, N.J. 1999. Complexities in the analysis of cryptic taxa

within the genus Anopheles. Parasitologia 41(1–3): 97–100.

Chandra, G., I. Bhattacharjee, and S. Chatterjee. 2010. A review of

Anopheles subpictus Grassi—a biological vector. Acta Tropica115: 142–154.

Chatterjee, S., and G. Chandra. 2000. Role of Anopheles subpictus as

a primary vector of malaria in an area in India. Japanese Journalof Tropical Medicine and Hygiene 28: 177–181.

Chaudhry, S., Neetu, S. Gupta, and J.S. Chhilar. 2005. Salivary

polytene chromosome mapping of Anopheles (Cellia) subpictusGrassi (Culicidae: Diptera). Genome 48: 241–246.

Chen, B., R.K. Butlin, and R.E. Harbach. 2003. Molecular phylog-

enetics of the Oriental members of the Myzomyia Series of

Anopheles subgenus Cellia (Diptera: Culicidae) inferred from

nuclear and mitochondrial DNA sequences. Systematic Ento-mology 28: 57–69.

Chhilar J.S. 2009. Molecular characterization of mosquito genome: A

study on emerging malaria vector Anopheles subpictus complex

(Culicidae: Diptera). PhD Thesis, Panjab University, Chandigarh.

Chhilar, J.S., and S. Chaudhry. 2005. Salivary polytene X-chromo-

some dynamics in Anopheles (Cellia) subpictus Grassi (Culic-

idae: Diptera). Journal of Cytology and Genetics, Bangalore 6:

57–63.

Collins, F.H., and S.M. Paskewitz. 1996. A review of the use of

ribosomal DNA (rDNA) to differentiate among cryptic Anoph-eles species. Insect Molecular Biology 5: 1–9.

Coluzzi, M. 1970. Sibling species in Anopheles and their importance

in malariology. Miscellaneous Publications of EntomologicalSociety of America 7: 63–77.

Cornel, A.J., C.H. Porter, and F.H. Collins. 1996. Polymerase chain

reaction species diagnostic assay for Anopheles quadrimaculatuscryptic species (Diptera: Culicidae) based on ribosomal DNA

ITS2 sequences. Journal of Medical Entomology 33: 109–116.

8 Proc Zool Soc (Jan-June 2012) 65(1):1–10

123

Dassanayake, R.S., Y.I.N.S. Gunawardene, and B.D.D.N.K. De Silva.

2008. ITS-2 secondary structures and phylogeny of Anophelesculicifacies species. Bioinformation 2: 456–460.

Favia, G., A. Lanfrancotti, L. Spanos, I. Siden-Kiamos, and C. Louis.

2001. Molecular characterization of ribosomal DNA polymor-

phisms discriminating among chromosomal forms of Anophelesgambiae s.s. Insect Molecular Biology 10: 19–23.

Felsenstein, J. 2004. PHYLIP (Phylogeny Inference Package) version3.6. Distributed by the author. Seattle, WA: Department of

Genome Sciences, University of Washington.

Foley, D.H., J.H. Bryan, D. Yeats, and A. Saul. 1998. Evolution and

systematics of Anopheles: Insights from a molecular phylogeny

of Australasian mosquitoes. Molecular Phylogenetics and Evo-lution 9: 262–275.

Foley, D.H., R.C. Wilkerson, R.D. Cooper, M.E. Volovsek, and J.H.

Bryan. 2007. A molecular phylogeny of Anopheles annulipes(Diptera: Culicidae) sensu lato: The most species-rich anophe-

line complex. Molecular Phylogenetics and Evolution 43:

283–297.

Fritz, G.N., J. Conn, A. Cockburn, and J. Seawright. 1994. Sequence

analysis of the ribosomal DNA internal transcribed spacer 2 from

populations of Anopheles nunegtovari (Diptera: Culicidae).

Molecular Biology and Evolution 11: 406–416.

Goswami, G., K. Raghavendra, N. Nanda, S.K. Gakhar, and S.K.

Subbarao. 2005. PCR-RFLP of mitochondrial cytochrome oxi-

dase subunit II and ITS2 of ribosomal DNA: Markers for the

identification of members of the Anopheles culicifacies complex

(Diptera: Culicidae). Acta Tropica 95: 92–99.

Green, C.A., B.A. Harrison, T. Klein, and V. Baimai. 1985. Cladistic

analysis of polytene chromosome rearrangements in Anopheline

mosquitoes, subgenus Cellia, series Neocellia. Canadian Journalof Genetics and Cytology 27: 123–133.

Harbach, R.E. 1994. A review of the internal classification of the

genus Anopheles (Diptera: Culicidae): The foundation for

comparative systematics and phylogenetic research. Bulletin ofEntomological Research 84: 331–342.

Harbach, R.E., and I.J. Kitching. 1998. Phylogeny and classification

of the Culicidae (Diptera). Systematic Entomology 23: 327–

370.

Harbach, R.E., and I.J. Kitching. 2005. Reconsideration of anopheline

mosquito phylogeny (Diptera: Culicidae: Anophelinae) based on

morphological data. Systematics and Biodiversity 3: 345–374.

Hubalek, Z., and J. Halouzka. 1999. West Nile Fever—a re-emerging

mosquito borne viral disease in Europe. Emerging InfectiousDiseases 5: 645–650.

Huong, N.T., P. Sonthayanon, A.J. Ketterman, and S. Panyim. 2001.

A rapid polymerase chain reaction based method for identifica-

tion of the Anopheles dirus sibling species. Southeast AsianJournal of Tropical Medicine and Public Health 32: 615–620.

Joseph, N., E. Krauskopf, M.I. Vera, and B. Michot. 1999. Ribosomal

internal transcribed spacer 2 (ITS2) exhibits a common core of

secondary structure in vertebrates and yeast. Nucleic AcidsResearch 27: 4533–4540.

Kampen, H., A. Sternberg, J. Proft, S. Bastian, F. Schaffner, W.A.

Maier, and H.M. Seitz. 2003. Polymerase chain reaction-based

differentiation of the mosquito sibling species Anopheles clav-iger s.s. and Anopheles petragnani (Diptera: Culicidae). Amer-ican Journal of Tropical Medicine and Hygiene 69: 195–199.

Kirti, J.S., and J. Kaur. 2004. Variations in ornamentation of wings

and palpi of Anopheles (Cellia) subpictus Grassi collected from

Northwest India. Journal of Vector Borne Diseases 41: 37–41.

Kitzmiller, J.B. 1976. Genetics, cytogenetics and evolution of

mosquitoes. Advances in Genetics 18: 315–433.

Knight, K.L., and A. Stone. 1977. A catalog of the mosquitoes of theworld (Diptera: Culicidae), vol. VI, 2nd ed. College Park, MD:

The Thomas Say Foundation. 387.

Krzywinski, J., and N.J. Besansky. 2003. Molecular systematics of

Anopheles: From subgenera to subpopulations. Annual Reviewsof Entomology 48: 111–139.

Krzywinski, J., R.C. Wilkerson, and N.J. Besansky. 2001. Evolution

of mitochondrial and ribosomal gene sequences in anophelinae

(Diptera: Culicidae): Implications for phylogeny reconstruction.

Molecular Phylogenetics and Evolution 18: 479–487.

Kulkarni, S.M. 1983. Detection of sporozoites in Anopheles subpictusin Bastar district, Madhya Pradesh. Indian Journal of Malari-ology 20: 159–160.

Linton, Y.M., R.E. Harbach, C.M. Seng, T.G. Anthony, and A.

Matusop. 2001. Morphological and molecular identity of

Anopheles (Cellia) sundaicus (Diptera: Culicidae), the nomino-

typical member of malaria vector species complex in south East

Asia. Systematic Entomology 26: 357–366.

Ma, Y., S. Li, and J. Xu. 2006. Molecular identification and

phylogeny of the Maculatus group of Anopheles mosquitoes

(Diptera: Culicidae) based on nuclear and mitochondrial DNA

sequences. Acta Tropica 99: 272–280.

Manonmani, A., H. Townson, T. Adeniran, P. Jambulingam, S. Sahu,

and T. Vijayakumar. 2001. rDNA-ITS2 polymerase chain

reaction assay for the sibling species of Anopheles fluviatilis.

Acta Tropica 78: 3–9.

Marinucci, M., R. Romi, P. Mancini, M. Di Luca, and C. Severini.

1999. Phylogenetic relationships of seven palearctic members of

the maculipennis complex inferred from ITS2 sequence analysis.

Insect Molecular Biology 8: 469–480.

Marrelli, M.T., R.S. Malafronte, C. Flores-Mendoza, R. Lourenco-de-

Oliveira, J.K. Kloetzel, and O. Marinotti. 1999. Sequence

analysis of the second internal transcribed spacer of ribosomal

DNA in Anopheles oswaldoi (Diptera: Culicidae). Journal ofMedical Entomology 36: 679–684.

Nagpal, B.N., and V.P. Sharma. 1995. Indian Anophelines. New

Delhi: Oxford and IBH Publishing Co. 416.

Nei, M. 1996. Phylogenetic analysis in molecular evolutionary

genetics. Annual Reviews of Genetics 30: 371–403.

Panicker, K.N., M. Geetharai, U.S. Bheema Rao, K. Viswam, and

M.V. Suryanarayana. 1981. Anopheles subpictus a vector of

malaria in coastal villages of south east India. Current Science50: 694–695.

Pape, T. 1992. Cladistic analyses of mosquito chromosome data in

Anopheles subgenus Cellia (Diptera: Culicidae). MosquitoSystematics 24: 1–11.

Paskewitz, S.M., D.M. Wesson, and F.H. Collins. 1993. The internal

transcribed spacers of ribosomal DNA in five members of the

Anopheles gambiae species complex. Insect Molecular Biology2: 247–297.

Philippe, H., F. Delsuc, H. Brinkmann, and N. Lartillot. 2005.

Phylogenomics. Annual Reviews of Ecology, Evolution, andSystematics 36: 541–562.

Phuc, H.K., A.J. Ball, L. Son, N.V. Hanh, N.D. Tu, N.G. Lien, A.

Verardi, and H. Townson. 2003. Multiplex PCR assay for

malaria vector Anopheles minimus and four related species in the

Myzomyia series from Southeast Asia. Medical and VeterinaryEntomology 17: 423–428.

Porter, C.H., and F.H. Collins. 1991. Species-diagnostic differences in

a ribosomal DNA internal transcribed spacer from the sibling

species Anopheles freeborni and Anopheles hermsi (Diptera:

Culicidae). American Journal of Tropical Medicine and Hygiene45: 271–279.

Rao, T.R. 1984. The Anophelines of India. New Delhi: Indian Council

of Medical Research. 518.

Retief, J.D. 2000. Phylogenetic analysis using PHYLIP. In Methods inmolecular biology, Vol 132: Bioinformatics methods and proto-cols, ed. S. Misener, and S.A. Krawetz, 243–258. Totowa, NJ:

Humuna Press.

Proc Zool Soc (Jan-June 2012) 65(1):1–10 9

123

Reuben, R., and S.G. Suguna. 1983. Morphological differences

between sibling species of the taxon Anopheles subpictus Grassi

in India, with notes on relationship with known forms. MosquitoNews 15: 117–126.

Sahu, S.S. 1998. Comparative susceptibility of Anopheles subpictusfrom fresh and brackish water areas to Plasmodium falciparuminfection. Acta Tropica 70: 1–7.

Sallum, M.A.M., T.R. Schultz, P.G. Foster, K. Aronstein, R.A. Wirtz,

and R.C. Wilkerson. 2002. Phylogeny of anophelinae (Diptera:

Culicidae) based on nuclear ribosomal and mitochondrial DNA

sequences. Systematic Entomology 27: 361–382.

Sambrook, J., E.F. Fritsch, and T. Maniatis. 1989. Molecular cloning:A laboratory manual, 2nd ed. Cold Spring Harbor, NY: Cold

Spring Harbor Laboratory Press.

Sanderson, M.J., and H. Bradley Shaffer. 2002. Troubleshooting

molecular phylogenetic analyses. Annual Reviews of Ecologyand Systematics 33: 49–72.

Schultz, J., S. Maisel, D. Gerlach, T. Muller, and M. Wolf. 2005. A

common core of secondary structure of the internal transcribed

spacer 2 (ITS2) throughout the Eukaryota. RNA 11: 361–364.

Subbarao, S.K., and V.P. Sharma. 1997. Anopheline species com-

plexes and malaria control. Indian Journal of Medical Research106: 164–173.

Suguna, S.G. 1982. Cytological and morphological evidence for

sibling species in Anopheles subpictus. Journal of Communica-ble Diseases 14: 1–8.

Suguna, S.G., K.G. Rathinam, A.R. Rajavel, and V. Dhanda. 1994.

Morphological and chromosomal descriptions of new species in

the Anopheles subpictus complex. Medical and VeterinaryEntomology 8: 88–94.

Surendran, S.N., O.P. Singh, P.J. Jude, and R. Ramasamy. 2010.

Genetic evidence for malaria vectors of the Anopheles sundaicuscomplex in Sri Lanka with morphological characteristics attrib-

uted to Anopheles subpictus species B. Malaria Journal 9: 343.

Tamura, K., J. Dudley, M. Nei, and S. Kumar. 2007. MEGA4:

Molecular Evolutionary Genetics Analysis (MEGA) software

version 4.0. Molecular Biology and Evolution 24: 1596–1599.

Wattal, B.L., and N.L. Kalra. 1967. Region-wise pictorial keys to the

female Indian Anopheles. Bulletin of the National Society ofIndia for Malaria and other Mosquito-Borne Diseases 9:

85–138.

WHO. 1999. Vectors in east Timor and malaria control activities. In

Report of preliminary TB and malaria assessment mission toDili, East Timor.

Xu, X., J. Xu, and F. Qu. 1998. A diagnostic polymerase chain

reaction assay for species A and D of the Anopheles dirus(Diptera: Culicidae) species complex based on ribosomal DNA

second internal transcribed spacer sequence. Journal of Amer-ican Mosquito Control Association 14: 385–389.

10 Proc Zool Soc (Jan-June 2012) 65(1):1–10

123