**Meet the editor**

Sylvie Manguin is a full-time Research Professor at the Institute of Research for Development (IRD), based at University of Montpellier. She is a leading medical entomologist and academician researcher whose main interest concerns mosquitoes involved in the transmission of pathogenic agents, especially Anopheles vectors of malaria agents. She has developed studies on

these mosquitoes from three continents (Asia, Africa, Americas) including species identification, population genetics, phylogenetic, vectorial capacities, spatial surveillance, midgut microbiota, immunological markers and vector control. She is the author of 75 indexed publications, four book chapters, and three books including "Biodiversity of malaria in the World" (John Libbey Ed.) that won the award of the International Festival of Medical Education Book (EDIMED). She is a member of the Editorial Boards of the Malaria Journal and Acta Tropica and serves as reviewer in several international institutions and more than 20 scientific journals.

## Contents



Gregory C. Lanzaro and Yoosook Lee



#### **Section 4 Pathogen Transmission and Influencing Factors 485**

Chapter 7 **Advances and Perspectives in the Study of the Malaria Mosquito Anopheles funestus 197**

*Anopheles funestus*

**Vectors in Africa 221**

**Southeast Asia 273**

**Southwest Pacific 357**

**Section 3 Ecology and Spatial Surveillance 395**

**Multidisciplinary Challenge 447**

Robert D. Cooper

Chapter 13 **Ecology of Larval Habitats 397**

*Anopheles*

Marc Coosemans

Roberts

Vas Dev and Vinod P. Sharma

Chapter 10 **Vector Biology and Malaria Transmission in**

**in India 239**

**VI** Contents

Ibrahima Dia, Moussa Wamdaogo Guelbeogo and Diego Ayala

*Anopheles nili Anopheles moucheti*

Wannapa Suwonkerd, Wanapa Ritthison, Chung Thuy Ngo, Krajana Tainchum, Michael J. Bangs and Theeraphap Chareonviriyaphap

Nigel W. Beebe, Tanya L. Russell, Thomas R. Burkot, Neil F. Lobo and

Eliška Rejmánková, John Grieco, Nicole Achee and Donald R.

Valérie Obsomer, Nicolas Titeux, Christelle Vancustem, Grégory Duveiller, Jean-François Pekel, Steve Connor, Pietro Ceccato and

Chapter 14 **From Anopheles to Spatial Surveillance: A Roadmap Through a**

Chapter 8 **Highlights on Anopheles nili and Anopheles moucheti, Malaria**

Christophe Antonio-Nkondjio and Frédéric Simard

Chapter 11 **Understanding Anopheles Diversity in Southeast Asia and Its**

Chapter 12 **The Systematics and Bionomics of Malaria Vectors in the**

Katy Morgan, Pradya Somboon and Catherine Walton

**Applications for Malaria Control 327**

*Anopheles*

Chapter 9 **The Dominant Mosquito Vectors of Human Malaria**

	- **Section 5 Vector Control: Current Situation, New Approaches and Perspectives 577**

## Chapter 23 **New Salivary Biomarkers of Human Exposure to Malaria Vector Bites 755**

Papa M. Drame, Anne Poinsignon, Alexandra Marie, Herbert Noukpo, Souleymane Doucoure, Sylvie Cornelie and Franck Remoue

## Chapter 24 **Transgenic Mosquitoes for Malaria Control: From the Bench to the Public Opinion Survey 797**

Christophe Boëte and Uli Beisel

## Preface

Chapter 23 **New Salivary Biomarkers of Human Exposure to Malaria**

Papa M. Drame, Anne Poinsignon, Alexandra Marie, Herbert Noukpo, Souleymane Doucoure, Sylvie Cornelie and Franck

Chapter 24 **Transgenic Mosquitoes for Malaria Control: From the Bench to**

**the Public Opinion Survey 797** Christophe Boëte and Uli Beisel

**Vector Bites 755**

Remoue

**VIII** Contents

First of all I would like to thank Sylvie Manguin, Editor of this book, who compiled 24 chap‐ ters that present current knowledge on malaria vector taxa. By asking me to preface this book, Sylvie got me out of my "bubble of Auvergne Region" and reconnected me with a scientific community that I had indeed never abandoned. It is with great pleasure that I found in this book the contributions of my former students and friends.

At the end of the second millennium, I had more or less put my pen down while the threat of global warming posed a major concern for the development and extension of vectorborne diseases. Particularly pessimistic forecasts predicted an extension of malaria up to the Polar Circle. However, no geographic expansion of malaria has been noticed in the last 20 years [1]. At most, the disease has reappeared on the Korean Peninsula where it was eradi‐ cated in the 1950s [2, 3]. Also, no particular invasion of *Anopheles* species has been observed as opposed to the global invasion of *Aedes albopictus* [4].

After the failure of the World Programme of Malaria Eradication (1950), WHO (World Health Organization) proposed at the Conference of Amsterdam (1992) a new strategy based on the treatment of all clinical cases using all chemo-therapeutic compounds, in par‐ ticular the artemisinin-based combination therapy (ACT). Vector control was a principal component of prevention. The use of insecticide-treated nets (ITN) has been shown to be effective in all epidemiological situations, and the pyrethroids used to impregnate the nets (permethrin, deltamethrin, lambdacyhalothrin, etc), besides protecting sleepers, has a bene‐ ficial impact on all members of communities where these nets are used [5, 6].

In last 20 years, manufacturers produced long-lasting insecticidal nets (LLIN) using fabrics that retain insecticide activity from three to five years (even after more than 15 washes). These LLINs are well accepted by users, and more than 24 million nets have been distribut‐ ed in the Afrotropical Region alone. In 1992, it was expected that implementation of this new strategy would initially reduce malaria mortality by 50% [6]. However, accurate data on malaria deaths is very difficult to obtain; this mainly rural disease often eludes official statistics and the results of different studies vary widely depending on the sources. The most recent estimates provided by Murray et al. in 2012 [7] give a more nuanced trend with the malaria mortality burden being larger than previously estimated, especially in adults. This study estimated that in 2010 malaria was the cause of 1.24 million deaths compared to 655,000 deaths reported by WHO, and in the Afrotropical Region infant mortality (children < 5 years old) due to malaria was estimated to be 24% *versus* 16% based on the WHO malaria report estimates [8]. These figures, although imprecise, provide a current estimate of the im‐ pact of malaria worldwide, which falls far short of the expected results despite the enor‐

mous financial expenditures of the WHO, charitable organizations, foundations and national initiatives.

Currently, the spectrum of resistance to many, if not the majority, of insecticides continues to pose a serious threat to all control programs, and alternate methods of control are of very limited efficacy [9]. Larval control by insecticides or insect growth regulators (IGR) is limit‐ ed to specific habitats, such as the oases of Oman. Hopes are now based on genetic control by transgenic mosquitoes. Research underway for more than 20 years has not produced a means of controlling malaria on a continental scale such as Africa where it endures without a solution for sustained control. We are still left with expectation.

Finally, one cannot ignore the considerable work on the systematics of *Anopheles* mosqui‐ toes. In addition to the creation of the subgenus *Baimaia* by Harbach, Rattanarithikul & Har‐ rison, many new species have been described or are waiting to be described [10, 11], especially in Asia where the majority of the vectors belong to species complexes [4]. South‐ east Asia with the *Anopheles dirus* complex and New Guinea with the *Anopheles farauti* com‐ plex, both comprised of eight species, represent 'hot spots' of *Anopheles* biodiversity. Species complexes include vectors and non-vectors and the identification of the vector species poses a real problem that can be solved by the new techniques apparent in the book.

This book, describing new insights and innovative approaches to the study of malaria vec‐ tors, contributes to a passionate aim of society – the eradication of malaria as a cause of mor‐ bidity and mortality in the poorest populations of the world.

> **Prof. Jean Mouchet** Emeritus Research Professor at IRD Langeac, France

#### **References**

[1] Reiter P. Global warming and malaria: knowing the horse before hitching the cart. Malar‐ ia Journal 2008;7 Suppl 1:S3.

[2] Manguin S, Carnevale P, Mouchet J, Coosemans M, Julvez J, Richard-Lenoble D, et al. Biodiversity of malaria in the world. Paris, France: John Libbey Eurotext; 2008.

[3] Mouchet J, Carnevale P, Coosemans M, Julvez J, Manguin S, Richard-Lenoble D, et al. Biodiversité du paludisme dans le monde. Paris, France: John Libbey Eurotext; 2004.

[4] Manguin S, Boëte C. Global impact of mosquito biodiversity, human vector-borne diseas‐ es and environmental change: InTech; 2011.

[5] Carnevale P, Robert V, Boudin C, Halna JM, Pazart L, Gazin P, et al. La lutte contre le paludisme par des moustiquaires imprégnées au Burkina Faso. Bull Soc Pathol Exot Filiales 1988;81:832-846.

[6] Lengeler C, Cattani JA, De Savigny D. Net Gain. A new method for the preventing ma‐ laria deaths. Geneva: WHO; 1996.

[7] Murray CJ, Rosenfeld LC, Lim SS, Andrews KG, Foreman KJ, Haring D, et al. Global malaria mortality between 1980 and 2010: a systematic analysis. Lancet 2012;379:413-431.

[8] WHO. World malaria report. In: WHO, editor. WHO global malaria program. Geneva: WHO; 2011. p. 278.

[9] malERA Consultative Group. A research agenda for malaria eradication: vector control. PLoS Med 2011;8:e1000401.

mous financial expenditures of the WHO, charitable organizations, foundations and

Currently, the spectrum of resistance to many, if not the majority, of insecticides continues to pose a serious threat to all control programs, and alternate methods of control are of very limited efficacy [9]. Larval control by insecticides or insect growth regulators (IGR) is limit‐ ed to specific habitats, such as the oases of Oman. Hopes are now based on genetic control by transgenic mosquitoes. Research underway for more than 20 years has not produced a means of controlling malaria on a continental scale such as Africa where it endures without

Finally, one cannot ignore the considerable work on the systematics of *Anopheles* mosqui‐ toes. In addition to the creation of the subgenus *Baimaia* by Harbach, Rattanarithikul & Har‐ rison, many new species have been described or are waiting to be described [10, 11], especially in Asia where the majority of the vectors belong to species complexes [4]. South‐ east Asia with the *Anopheles dirus* complex and New Guinea with the *Anopheles farauti* com‐ plex, both comprised of eight species, represent 'hot spots' of *Anopheles* biodiversity. Species complexes include vectors and non-vectors and the identification of the vector species poses

This book, describing new insights and innovative approaches to the study of malaria vec‐ tors, contributes to a passionate aim of society – the eradication of malaria as a cause of mor‐

[1] Reiter P. Global warming and malaria: knowing the horse before hitching the cart. Malar‐

[2] Manguin S, Carnevale P, Mouchet J, Coosemans M, Julvez J, Richard-Lenoble D, et al.

[3] Mouchet J, Carnevale P, Coosemans M, Julvez J, Manguin S, Richard-Lenoble D, et al.

[4] Manguin S, Boëte C. Global impact of mosquito biodiversity, human vector-borne diseas‐

[5] Carnevale P, Robert V, Boudin C, Halna JM, Pazart L, Gazin P, et al. La lutte contre le paludisme par des moustiquaires imprégnées au Burkina Faso. Bull Soc Pathol Exot Filiales

[6] Lengeler C, Cattani JA, De Savigny D. Net Gain. A new method for the preventing ma‐

[7] Murray CJ, Rosenfeld LC, Lim SS, Andrews KG, Foreman KJ, Haring D, et al. Global malaria mortality between 1980 and 2010: a systematic analysis. Lancet 2012;379:413-431. [8] WHO. World malaria report. In: WHO, editor. WHO global malaria program. Geneva:

Biodiversity of malaria in the world. Paris, France: John Libbey Eurotext; 2008.

Biodiversité du paludisme dans le monde. Paris, France: John Libbey Eurotext; 2004.

**Prof. Jean Mouchet**

Langeac, France

Emeritus Research Professor at IRD

a solution for sustained control. We are still left with expectation.

bidity and mortality in the poorest populations of the world.

a real problem that can be solved by the new techniques apparent in the book.

national initiatives.

X Preface

**References**

1988;81:832-846.

WHO; 2011. p. 278.

ia Journal 2008;7 Suppl 1:S3.

es and environmental change: InTech; 2011.

laria deaths. Geneva: WHO; 1996.

[10] Harbach RE. The classification of genus *Anopheles* (Diptera: Culicidae): a working hy‐ pothesis of phylogenetic relationships. Bull Entomol Res 2004;94:537-553.

[11] Harbach RE, Rattanarithikul R, Harrison BA. *Baimaia*, a new subgenus for *Anopheles kyondawensis* Abraham, a unique crabhole-breeding anopheline in Southeast Asia. The Pro‐ ceedings of the Entomological Society of Washington 2005;107.

#### **Preface from the Editor**

In a global public health context, the genus *Anopheles* is by far the most important group of pathogen-carrying mosquitoes due to their exclusive involvement in the transmission of hu‐ man malaria parasites. To properly control malaria, an entirely preventable and treatable disease, the current recommended interventions advocated by WHO [1] include vector con‐ trol through the use of insecticide-treated nets (ITNs), periodic indoor residual spraying (IRS) and, in some specific settings, larval control. During the past decade, coverage with vector control interventions have increased substantially in sub-Saharan Africa; for example, reaching 53% of households with at least one ITN in 2012. However, WHO has also ob‐ served that *"due to fewer deliveries of ITNs and increasing mosquito resistance to insecticides, re‐ cent successes in malaria vector control may be jeopardized."* [1]. As vector control is a very effective means of malaria control, a better understanding of *Anopheles* populations is a key element for reaching the goal of malaria elimination in the future [2].

A large amount of scientific knowledge and technical advances concerning these mosquitoes has accumulated over the past century, and in recent decades the advent of novel technolo‐ gies have accelerated the acquisition of new information. In fact, the current trend of re‐ search and new findings as a consequence of the renewed emphasis for controlling malaria using vector control is rapidly expanding our understanding of *Anopheles* mosquitoes. The 24 chapters of this book present some of the latest research on important malaria vectors using innovative approaches supported by state-of-the-art methodologies covering a wide array of study disciplines on the biology, genetics, distribution, pathogen transmission, and the application of these findings in the improvement of current vector control strategies.

This book is divided into five sections. Section 1: Focuses on the reliable identification and classification of certain species, an area that has been fraught with past difficulties for accu‐ rately differentiating the individual sibling species placed within taxonomic complexes. The precise identification of a species must be linked to their specific role and importance in the transmission of malaria agents that can dramatically differ from their morphologically indis‐ tinguishable sibling species. Section 2: Provides up-to-date information on the genetic diver‐ sity, bionomics and distribution of the dominant vector species of Latin America, Africa, Asia, and Southwest Pacific as presented in nine chapters. Section 3: Presents a better under‐ standing of environmental aspects linked to larval habitat ecology and spatial surveillance of *Anopheles* vectors having found increasing utility in the last decade. Section 4: Pathogen transmission is presented with a focus on *Plasmodium knowlesi,* an emerging public health problem in Southeast Asia, along with transmission influencing factors such as thermoregu‐ lation during blood feeding and the role of mosquito midgut microbiota as possible means to control disease by transmission blocking approach. Section 5: Reviews the current status on insecticide resistance, innovative approaches to vector control, and new tools to evaluate control efficacy based on *Anopheles* saliva biomarkers and perspectives for using transgenic mosquitoes.

A total of 71 authors, from 20 countries on five continents (Africa, Asia, Australia, America, and Europe), with internationally recognized expertise, have generously participated in this book and I am extremely grateful to all for their time and energy to contribute to new and innovative topics on *Anopheles*. The originality of this book, published as 'open access' by InTech, is to offer a detailed description and analysis of new concepts, paradigms and inno‐

vative approaches on the understanding of *Anopheles* mosquitoes and the development of better weapons to control the vector.

I am also profoundly grateful to Professor Jean Mouchet, my mentor and the one of genera‐ tions of medical entomologists, who kindly accepted to write a preface for this book. I also thank Ms. Dragana Manestar for her valuable support and assistance in publishing this book. This publication should benefit not only medical entomologists, but also students, sci‐ entists, public health managers, and decision makers interested in malaria and its vectors.

**Sylvie Manguin, PhD**

Institute of Research for Development (IRD) Montpellier, France

#### **References**

**Preface from the Editor**

XII Preface

mosquitoes.

In a global public health context, the genus *Anopheles* is by far the most important group of pathogen-carrying mosquitoes due to their exclusive involvement in the transmission of hu‐ man malaria parasites. To properly control malaria, an entirely preventable and treatable disease, the current recommended interventions advocated by WHO [1] include vector con‐ trol through the use of insecticide-treated nets (ITNs), periodic indoor residual spraying (IRS) and, in some specific settings, larval control. During the past decade, coverage with vector control interventions have increased substantially in sub-Saharan Africa; for example, reaching 53% of households with at least one ITN in 2012. However, WHO has also ob‐ served that *"due to fewer deliveries of ITNs and increasing mosquito resistance to insecticides, re‐ cent successes in malaria vector control may be jeopardized."* [1]. As vector control is a very effective means of malaria control, a better understanding of *Anopheles* populations is a key

A large amount of scientific knowledge and technical advances concerning these mosquitoes has accumulated over the past century, and in recent decades the advent of novel technolo‐ gies have accelerated the acquisition of new information. In fact, the current trend of re‐ search and new findings as a consequence of the renewed emphasis for controlling malaria using vector control is rapidly expanding our understanding of *Anopheles* mosquitoes. The 24 chapters of this book present some of the latest research on important malaria vectors using innovative approaches supported by state-of-the-art methodologies covering a wide array of study disciplines on the biology, genetics, distribution, pathogen transmission, and the application of these findings in the improvement of current vector control strategies. This book is divided into five sections. Section 1: Focuses on the reliable identification and classification of certain species, an area that has been fraught with past difficulties for accu‐ rately differentiating the individual sibling species placed within taxonomic complexes. The precise identification of a species must be linked to their specific role and importance in the transmission of malaria agents that can dramatically differ from their morphologically indis‐ tinguishable sibling species. Section 2: Provides up-to-date information on the genetic diver‐ sity, bionomics and distribution of the dominant vector species of Latin America, Africa, Asia, and Southwest Pacific as presented in nine chapters. Section 3: Presents a better under‐ standing of environmental aspects linked to larval habitat ecology and spatial surveillance of *Anopheles* vectors having found increasing utility in the last decade. Section 4: Pathogen transmission is presented with a focus on *Plasmodium knowlesi,* an emerging public health problem in Southeast Asia, along with transmission influencing factors such as thermoregu‐ lation during blood feeding and the role of mosquito midgut microbiota as possible means to control disease by transmission blocking approach. Section 5: Reviews the current status on insecticide resistance, innovative approaches to vector control, and new tools to evaluate control efficacy based on *Anopheles* saliva biomarkers and perspectives for using transgenic

A total of 71 authors, from 20 countries on five continents (Africa, Asia, Australia, America, and Europe), with internationally recognized expertise, have generously participated in this book and I am extremely grateful to all for their time and energy to contribute to new and innovative topics on *Anopheles*. The originality of this book, published as 'open access' by InTech, is to offer a detailed description and analysis of new concepts, paradigms and inno‐

element for reaching the goal of malaria elimination in the future [2].

[1] WHO. WHO malaria report. In: WHO, editor. WHO global malaria program. Geneva: WHO; 2012. p. 260.

[2] malERA Consultative Group. A research agenda for malaria eradication: vector control. PLoS Med 2011;8:e1000401.

**Species Identification and Phylogeny of Anopheles**

## **Chapter 1**

## **The Phylogeny and Classification of** *Anopheles*

Ralph E. Harbach

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/54695

## **1. Introduction**

*Anopheles* was introduced as a genus of mosquitoes in 1818 by Johann Wilhelm Meigen [1], a German entomologist famous for his revolutionary studies of Diptera. Little was done on the taxonomy of *Anopheles* until the discovery during the last two decades of the 19th century that mosquitoes transmit microfilariae and malarial protozoa, which initiated a drive to collect, name and classify these insects. In 1898, the Royal Society and the Rt. Hon. Joseph Chamber‐ lain, Secretary of State for the Colonies of Britain, appointed a Committee to supervise the investigation of malaria. On 6 December 1898, Mr. Chamberlain directed the Colonies to collect and send mosquitoes to the British Museum (Natural History) (Figure 1), and in 1899 the Committee appointed Frederick V. Theobald to prepare a monograph on the mosquitoes of the world, which was published in five volumes between 1901 and 1910 [2‒6]. As a conse‐ quence, many new generic names were introduced in an effort to classify numerous new mosquito species into seemingly natural groups. Theobald proposed 18 genera for species of *Anopheles* based on the distribution and shape of scales on the thorax and abdomen. Four of these proposed genera, *Cellia*, *Kerteszia*, *Nyssorhynchus* and *Stethomyia*, are currently recognized as subgenera of *Anopheles* and the other 14 are regarded as synonyms of one or other of subgenera *Anopheles*, *Cellia* or *Nyssorhynchus*. Theobald, however, was not the only person to propose generic names for species of *Anopheles*. During the first three decades of the 20th century, 37 genera (including the 18 recognized by Theobald) were established for species of *Anopheles* [7].

As additional new species were discovered, it became increasingly apparent that Theobald's system of classification was neither practical nor natural. Frederick Knab in North America, one of the early critics of Theobald's classification, stated that "the subject was made needlessly difficult by hasty work and by the sub-division of the old genus *Anopheles* into numerous illdefined and fancifully differentiated genera. The intricacies of this 'system,' unwarranted from both a scientific and practical standpoint, even the trained entomologist could not tread with

© 2013 Harbach; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Harbach; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

safety, and to others it could be no less than hopeless or disastrous" [8]. Consequently, during the two decades following the completion of Theobald's monograph in 1910, significant changes were made toward a much more conservative system of classification, culminating in the reduction of 38 genus-group names (including *Anopheles*) to the recognition of the single genus *Anopheles*.

The current subgeneric classification of *Anopheles* is based primarily on the number and positions of specialized setae on the gonocoxites of the male genitalia (Figure 2), and this basis of classification has been accepted since it was introduced by Sir (Samuel) Rickard Christo‐ phers in 1915 [9]. Christophers proposed three generic subdivisions, which F.M. Edwards [10] and Francis Metcalf Root [11] formally recognized as subgenera *Anopheles*, *Myzomyia* (=*Cellia*) and *Nyssorhynchus*. Edwards adopted this system and added subgenus *Stethomyia* in his classical treatise on family Culicidae published in 1932 [12]. This system recognized *Kerteszia* as an informal group within subgenus *Nyssorhynchus*. *Kerteszia* was elevated to subgeneric status by W.H.W. Komp [13]. Subgenus *Lophopodomyia* was proposed by P.C.A. Antunes in 1937 [14] and subgenus *Baimaia* was introduced by Ralph E. Harbach and his colleagues in 2005 [15].

Genus *Anopheles* currently includes 465 formally named species that are disproportionately divided between seven subgenera: *Anopheles* (cosmopolitan, 182 species), *Baimaia* (Oriental, one species), *Cellia* (Old World, 220 species), *Kerteszia* (Neotropical, 12 species), *Lophopodo‐ myia* (Neotropical, six species), *Nyssorhynchus* (Neotropical, 39 species) and *Stethomyia* (Neotropical, five species) [16]. Four of the subgenera, *Anopheles*, *Cellia*, *Kerteszia* and *Nysso‐ rhynchus*, include the species that transmit human malarial parasites. Most vector species of *Anopheles* have been found to comprise complexes of sibling species.

## **2. Classification of genus** *Anopheles*

The aim of classification is to group and categorize biological entities that share some unifying characteristics. Classification has been defined by Ernst Mayr & W.J. Bock [17] as "The arrangement of similar entities (objects) in a hierarchical series of nested classes, in which each more inclusive higher-level class is subdivided comprehensively into less inclusive classes at the next lower level." These classes (groups) are known as **taxa** (singular: **taxon**). The level of a taxon in a hierarchical classification is referred to as a **taxonomic rank** or **category**. Ideally, taxonomic categories should denote equivalent phylogenetic rank; however, in practice they are basically subjective groupings of subordinate taxa that are presumed to represent mono‐ phyletic groups of species that are assigned to taxonomic ranks based on shared morphological and biological characteristics that are not a measure of phylogenetic equivalence. For this reason, the taxonomic categories of genus *Anopheles*, including the formal rank of subgenus, should not be considered to represent equivalent phylogenetic ranks.

safety, and to others it could be no less than hopeless or disastrous" [8]. Consequently, during the two decades following the completion of Theobald's monograph in 1910, significant changes were made toward a much more conservative system of classification, culminating in the reduction of 38 genus-group names (including *Anopheles*) to the recognition of the single

The current subgeneric classification of *Anopheles* is based primarily on the number and positions of specialized setae on the gonocoxites of the male genitalia (Figure 2), and this basis of classification has been accepted since it was introduced by Sir (Samuel) Rickard Christo‐ phers in 1915 [9]. Christophers proposed three generic subdivisions, which F.M. Edwards [10] and Francis Metcalf Root [11] formally recognized as subgenera *Anopheles*, *Myzomyia* (=*Cellia*) and *Nyssorhynchus*. Edwards adopted this system and added subgenus *Stethomyia* in his classical treatise on family Culicidae published in 1932 [12]. This system recognized *Kerteszia* as an informal group within subgenus *Nyssorhynchus*. *Kerteszia* was elevated to subgeneric status by W.H.W. Komp [13]. Subgenus *Lophopodomyia* was proposed by P.C.A. Antunes in 1937 [14] and subgenus *Baimaia* was introduced by Ralph E. Harbach and his colleagues in

Genus *Anopheles* currently includes 465 formally named species that are disproportionately divided between seven subgenera: *Anopheles* (cosmopolitan, 182 species), *Baimaia* (Oriental, one species), *Cellia* (Old World, 220 species), *Kerteszia* (Neotropical, 12 species), *Lophopodo‐ myia* (Neotropical, six species), *Nyssorhynchus* (Neotropical, 39 species) and *Stethomyia* (Neotropical, five species) [16]. Four of the subgenera, *Anopheles*, *Cellia*, *Kerteszia* and *Nysso‐ rhynchus*, include the species that transmit human malarial parasites. Most vector species of

The aim of classification is to group and categorize biological entities that share some unifying characteristics. Classification has been defined by Ernst Mayr & W.J. Bock [17] as "The arrangement of similar entities (objects) in a hierarchical series of nested classes, in which each more inclusive higher-level class is subdivided comprehensively into less inclusive classes at the next lower level." These classes (groups) are known as **taxa** (singular: **taxon**). The level of a taxon in a hierarchical classification is referred to as a **taxonomic rank** or **category**. Ideally, taxonomic categories should denote equivalent phylogenetic rank; however, in practice they are basically subjective groupings of subordinate taxa that are presumed to represent mono‐ phyletic groups of species that are assigned to taxonomic ranks based on shared morphological and biological characteristics that are not a measure of phylogenetic equivalence. For this reason, the taxonomic categories of genus *Anopheles*, including the formal rank of subgenus,

*Anopheles* have been found to comprise complexes of sibling species.

should not be considered to represent equivalent phylogenetic ranks.

**2. Classification of genus** *Anopheles*

genus *Anopheles*.

4 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

2005 [15].

**Figure 1.** Letter issued from Downing Street on 6 December 1898 directing the British Colonies to collect and send mosquitoes to the British Museum (Natural History).

**Figure 2.** Subgenera of *Anopheles* ‒ specialized setae on the gonocoxites of the male genitalia (after Harbach & Kitch‐ ing [18]): A, *Anopheles*; B. *Baimaia*; C, *Cellia*; D, *Kerteszia*; E, *Lophopodomyia*; F, *Nyssorhynchus*; G, *Stethomyia*. as, ac‐ cessory setae; is, inner seta; ps, parabasal seta(e).

Infrasubgeneric categories (taxonomic ranks below subgenus) have no formal status under the *International Code of Zoological Nomenclature* [19]. They are convenience categories only, often based on superficial similarities that may not indicate natural relationships. The informal categories used in the classification of *Anopheles* include Sections, Series, Groups, Subgroups and Complexes (see Appendix 1).

Unlike formal taxonomic categories, which precede the name of the taxonomic unit, for instance family Culicidae, genus *Anopheles* and species *gambiae*, the names of informal taxonomic categories follow the name of the taxonomic unit, for example the Pyretophorus Series, Hyrcanus Group or Gambiae Complex, which are written in Roman (i.e. non-italic) script with the first letter capitalized. It should be stressed that both formal and informal taxonomic entities are conceptual constructs invented by taxonomists for the purpose of creating some order in the diversity of species. For example, the species *gambiae* and the Hyrcanus Group, which are human-conceived taxonomic concepts, cannot be observed as entities or visualized under a microscope.

The internal classification of genus *Anopheles* (between genus and species levels) is based primarily on the schemes proposed by Edwards [12], John A. Reid & Kenneth L. Knight [20], Alexis Grjebine [21], M.T. Gillies & Botha de Meillon [22], Reid [23], Michael E. Faran [24] and Kenneth J. Linthicum [25]. These schemes were reviewed, amalgamated and updated in 1994 [26] and updated again in 2004 and 2012 [27,16 respectively]. The three largest subgenera, i.e. *Anopheles*, *Cellia* and *Nyssorhynchus*, are divided into hierarchical systems of informal taxo‐ nomic categories (Appendix 1; examples shown in Figure 3). Subgenus *Anopheles* is divided into two Sections based on the shape of the pupal trumpet. The Laticorn Section was created for species with a wide funnel-shaped trumpet having the longest axis transverse to the stem, and the Angusticorn Section for species with a semi-tubular trumpet having the longest axis vertical more or less in line with the stem [20]. Subgenus *Nyssorhynchus* is divided into three Sections based on unique combinations of larval, pupal and adult characters [28]. Subgenus *Cellia* and the Sections of subgenera *Anopheles* and *Nyssorhynchus* are divided into Series, the larger Series are divided into species Groups, and some Groups are further divided into Subgroups and species Complexes. Most of the groupings at each level of classification are presumed to represent natural groups of species, thus implying phylogenetic relationships, but much additional basic taxonomic research is needed before the formal and informal taxa can be firmly established as monophyletic entities. The internal classification of the genus (subgenera and infrasubgeneric groups) is detailed in Appendix 1. An alphabetical list of all formally named, currently recognized species and their position in the classification is provided in Appendix 2. Similarly, all currently known sibling species complexes are listed in Appendix 3, and the unnamed and provisionally designated species of the complexes and their position in the classification are listed in Appendix 4.

## **3. Phylogeny of** *Anopheles*

**Figure 2.** Subgenera of *Anopheles* ‒ specialized setae on the gonocoxites of the male genitalia (after Harbach & Kitch‐ ing [18]): A, *Anopheles*; B. *Baimaia*; C, *Cellia*; D, *Kerteszia*; E, *Lophopodomyia*; F, *Nyssorhynchus*; G, *Stethomyia*. as, ac‐

Infrasubgeneric categories (taxonomic ranks below subgenus) have no formal status under the *International Code of Zoological Nomenclature* [19]. They are convenience categories only, often based on superficial similarities that may not indicate natural relationships. The informal categories used in the classification of *Anopheles* include Sections, Series, Groups, Subgroups

Unlike formal taxonomic categories, which precede the name of the taxonomic unit, for instance family Culicidae, genus *Anopheles* and species *gambiae*, the names of informal taxonomic categories follow the name of the taxonomic unit, for example the Pyretophorus Series, Hyrcanus Group or Gambiae Complex, which are written in Roman (i.e. non-italic) script with the first letter capitalized. It should be stressed that both formal and informal taxonomic entities are conceptual constructs invented by taxonomists for the purpose of creating some order in the diversity of species. For example, the species *gambiae* and the Hyrcanus Group, which are human-conceived taxonomic concepts, cannot be observed as

The internal classification of genus *Anopheles* (between genus and species levels) is based primarily on the schemes proposed by Edwards [12], John A. Reid & Kenneth L. Knight [20], Alexis Grjebine [21], M.T. Gillies & Botha de Meillon [22], Reid [23], Michael E. Faran [24] and Kenneth J. Linthicum [25]. These schemes were reviewed, amalgamated and updated in 1994 [26] and updated again in 2004 and 2012 [27,16 respectively]. The three largest subgenera, i.e. *Anopheles*, *Cellia* and *Nyssorhynchus*, are divided into hierarchical systems of informal taxo‐

cessory setae; is, inner seta; ps, parabasal seta(e).

6 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

and Complexes (see Appendix 1).

entities or visualized under a microscope.

*Anopheles* is undoubtedly the most studied and best known genus of mosquitoes, largely because of their great impact on human health. As vectors of causative agents of malaria and filariasis, *Anopheles* mosquitoes have affected the lives of more humans than any other insects. As a matter of fact, *Anopheles* is one of few groups of eukaryote organisms that have had an impact on human evolution ‒ the emergence of sickle cell anemia as a mode of resistance to malarial protozoa. As a result of more than a century of studies by medical entomologists, taxonomists and geneticists, 537 species of *Anopheles* are currently known and most have been formally named (87%) (Appendix 2), but until recently little work has been done to understand the evolution and phylogenetic relationships of these mosquitoes.

**Figure 3.** Hierarchical classification (from specific to general) of A. *Anopheles freeborni*, Freeborni Subgroup, Maculi‐ pennis Group, Anopheles Series, Angusticorn Section, Subgenus *Anopheles*; B. *Anopheles minimus*, Minimus Complex, Minimus Subgroup, Funestus Group, Myzomyia Series, Subgenus *Cellia*; C. *Anopheles albimanus*, Albimanus Series, Al‐ bimanus Section, Subgenus *Nyssorhynchus*.

The phylogenetic studies of anopheline mosquitoes conducted to date are summarized in Appendix 5. In view of the impact of malaria on human health, it is not surprising that most of these studies have dealt with species Groups, Subgroups and Complexes that include vectors of human malarial protozoa. It is obvious that the evolutionary relationships of malaria vectors and their closest allies have received more attention than other groups. However, none of these studies can be regarded as complete in terms of taxonomic coverage of any group, and the field of disease vector systematics presents many opportunities for further research. Phylogenetic patterns are used to interpret bionomic features such as differences in the nature of blood-feeding by adult females, feeding behavior and the occurrence of immature stages in aquatic habitats.

Mosquitoes probably evolved in the Jurassic [12,29,30] (146‒200 Mya)<sup>1</sup> , along with the early mammals, first birds and first flowering plants. Unfortunately, due to the paucity of mosquito fossils, there is no direct indication of the evolutionary history of anopheline mosquitoes. The second oldest fossil mosquito, *Paleoculicis minutus* [31] from the Late Cretaceous (66.0–100.5 Mya), has morphological features that indicate a closer affinity with culicine than anopheline mosquitoes, which suggests that this ancestral lineage is younger than the lineage that gave rise to subfamily Anophelinae. *Anopheles*(*Nyssorhynchus*?) *dominicanus*[32] and *An*. (?) *rottensis* [33] are the only fossil anopheline mosquitoes. The former is from the mid-Tertiary (about 15– 45 Mya) and the latter is from the Late Oligocene of Germany (approximately 25 Mya). If the anopheline mosquitoes are indeed ancestral to all other Culicidae [18,34], it would appear from available fossil evidence that extant groups may have evolved in the Cenozoic Era (<66.0 Mya). From divergence times based on sequence data for nuclear protein-coding genes and fossil calibration points, it appears that major mosquito lineages date to the Early Cretaceous (100.5– 145.0 Mya), and the ancestral lineage of anophelines may have appeared as early as the Jurassic (~145 Mya) [34].

*Anopheles* is the nominotypical genus of subfamily Anophelinae. In addition to *Anopheles* (cosmopolitan), the subfamily includes two other genera: *Bironella* (Australasian) and *Chagasia* (Neotropical). Cladistic analyses of morphological data and DNA sequences of various ribosomal, mitochondrial and nuclear genes strongly support the placement of *Chagasia* in an ancestral relationship to all other anophelines [18,34‒41].

In 2000, Sallum et al. [40] performed the first phylogenetic analysis of subfamily Anophelinae, based on morphological characters. The results indicated that genus *Anopheles* is paraphyletic because it included genus *Bironella*. Subgenera *Kerteszia*, *Nyssorhynchus*, *Cellia*, *Lophopodomyia* and *Stethomyia*, along with genus *Bironella*, were found to be monophyletic taxa dispersed among various Series and species Groups of subgenus *Anopheles*. The Christya Series of subgenus *Anopheles* was placed with *Kerteszia* + *Nyssorhynchus* and this clade was sister to *Cellia* + all other anophelines except *Chagasia*.

Two years later, Sallum et al. [41] conducted a molecular analysis of anopheline relationships based on ribosomal (18S, 28S) and mitochondrial (COI, COII) DNA sequences. The results of

<sup>1</sup> Geological ages of eras and periods follow the geological timescale determined by the International Commission on Stratigraphy (http://www.stratigraphy.org).

that study cannot be compared directly with the results of their earlier study [40] because significantly fewer taxa were included in the analyses. Nevertheless, the molecular data corroborated the paraphyly of genus *Anopheles* relative to *Bironella* and the sister-group relationship of *Kerteszia* and *Nyssorhynchus*, and supported the monophyly of the other subgenera and genus *Bironella*, which was reconstructed as the sister to *Lophopodomyia* rather than *Stethomyia*.

The phylogenetic studies of anopheline mosquitoes conducted to date are summarized in Appendix 5. In view of the impact of malaria on human health, it is not surprising that most of these studies have dealt with species Groups, Subgroups and Complexes that include vectors of human malarial protozoa. It is obvious that the evolutionary relationships of malaria vectors and their closest allies have received more attention than other groups. However, none of these studies can be regarded as complete in terms of taxonomic coverage of any group, and the field of disease vector systematics presents many opportunities for further research. Phylogenetic patterns are used to interpret bionomic features such as differences in the nature of blood-feeding by adult females, feeding behavior and the occurrence of immature stages in

mammals, first birds and first flowering plants. Unfortunately, due to the paucity of mosquito fossils, there is no direct indication of the evolutionary history of anopheline mosquitoes. The second oldest fossil mosquito, *Paleoculicis minutus* [31] from the Late Cretaceous (66.0–100.5 Mya), has morphological features that indicate a closer affinity with culicine than anopheline mosquitoes, which suggests that this ancestral lineage is younger than the lineage that gave rise to subfamily Anophelinae. *Anopheles*(*Nyssorhynchus*?) *dominicanus*[32] and *An*. (?) *rottensis* [33] are the only fossil anopheline mosquitoes. The former is from the mid-Tertiary (about 15– 45 Mya) and the latter is from the Late Oligocene of Germany (approximately 25 Mya). If the anopheline mosquitoes are indeed ancestral to all other Culicidae [18,34], it would appear from available fossil evidence that extant groups may have evolved in the Cenozoic Era (<66.0 Mya). From divergence times based on sequence data for nuclear protein-coding genes and fossil calibration points, it appears that major mosquito lineages date to the Early Cretaceous (100.5– 145.0 Mya), and the ancestral lineage of anophelines may have appeared as early as the Jurassic

*Anopheles* is the nominotypical genus of subfamily Anophelinae. In addition to *Anopheles* (cosmopolitan), the subfamily includes two other genera: *Bironella* (Australasian) and *Chagasia* (Neotropical). Cladistic analyses of morphological data and DNA sequences of various ribosomal, mitochondrial and nuclear genes strongly support the placement of *Chagasia* in an

In 2000, Sallum et al. [40] performed the first phylogenetic analysis of subfamily Anophelinae, based on morphological characters. The results indicated that genus *Anopheles* is paraphyletic because it included genus *Bironella*. Subgenera *Kerteszia*, *Nyssorhynchus*, *Cellia*, *Lophopodomyia* and *Stethomyia*, along with genus *Bironella*, were found to be monophyletic taxa dispersed among various Series and species Groups of subgenus *Anopheles*. The Christya Series of subgenus *Anopheles* was placed with *Kerteszia* + *Nyssorhynchus* and this clade was sister to *Cellia*

Two years later, Sallum et al. [41] conducted a molecular analysis of anopheline relationships based on ribosomal (18S, 28S) and mitochondrial (COI, COII) DNA sequences. The results of

1 Geological ages of eras and periods follow the geological timescale determined by the International Commission on

, along with the early

Mosquitoes probably evolved in the Jurassic [12,29,30] (146‒200 Mya)<sup>1</sup>

ancestral relationship to all other anophelines [18,34‒41].

+ all other anophelines except *Chagasia*.

Stratigraphy (http://www.stratigraphy.org).

aquatic habitats.

8 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

(~145 Mya) [34].

In 2005, Harbach & Kitching [36] revised and expanded the phylogenetic analysis of Sallum et al. [40], with special consideration of the specialized setae of the male gonocoxites (Figure 2) that diagnose the subgenera. Parsimony analysis of the data set under implied weighting supported the monophyly of subgenera *Cellia*, *Kerteszia* and *Nyssorhynchus*, and the sister relationship of *Kerteszia* + *Nyssorhynchus*. Subgenus *Anopheles* was recovered as a polyphyletic lineage basal to a monophyletic clade consisting of *Kerteszia* + *Nyssorhynchus* and *Cellia* in a sister-group relationship. *Bironella*, *Lophopodomyia* and *Stethomyia* were firmly nested within subgenus *Anopheles*, which would be paraphyletic even if these taxa were subsumed within it. Subgenus *Baimaia*, represented by *An. kyondawensis*, was supported as the sister of *Bironel‐ la* + all other *Anopheles*. *Bironella* and *Stethomyia*, contrary to the earlier study of Sallum et al. [40], were also supported as monophyletic clades separate from subgenus *Anopheles*. The preferred cladogram of Harbach & Kitching (Figures 4 and 5) is taken here to represent the best available estimate of anopheline phylogeny and evolutionary relationships because it is based on a greater number of taxonomic groups and homologous characters than all other hypotheses published to date.

A later analysis of subgenus *Anopheles* by Collucci & Sallum [42] included 38 species repre‐ senting the same Series (6) and species Groups (15) of the subgenus that were included in the study of Sallum et al. [40]. The data were analyzed using successive approximations character weighting (SACW) and implied weighting (IW). Most of the relationships between members of the subgenus were either moderately or poorly supported. The Laticorn Section was recovered as a monophyletic clade in the IW analysis, suggesting that the laticorn development of the pupal trumpet is a derived condition for subgenus *Anopheles*. In the SACW analyses, members of the group comprised a paraphyletic lineage relative to the Cycloleppteron Series. The Angusticorn Section was recovered as a polyphyletic assemblage in both analyses. These results are contradicted by those of Sallum et al. [40] and Harbach & Kitching [36] who found that neither section is monophyletic. Below the section level of classification, only the Lopho‐ scelomyia and Arribalzagia Series were recovered as monophyletic assemblages. The Myzo‐ rhynchus Series was paraphyletic relative to the Cycloleppteron, Christya and Arribalzagia Series, and the Anopheles Series was polyphyletic. Surprisingly, the two species of the Cycloleppteron Series included in the analyses were not grouped together, suggesting that the series is not monophyletic. In contrast, the Arribalzagia, Christya, Cycloleppteron, Lophosce‐ lomyia and Myzorhynchus Series were recovered as monophyletic assemblages in the IW analysis of Harbach & Kitching (Figure 4). Furthermore, with the removal of subgenus *Baimaia*, the remaining species of the Anopheles Series included in their analysis also formed a monophyletic group. With the exception of the Pseudopunctipennis Group, all the species groups represented in the analysis of Collucci & Sallum (Aitkenii, Albotaeniatus, Culiciformis, Hyrcanus, Plumbeus, Umbrosus Groups) were recovered as monophyletic assemblages with moderate to strong support [the Pseudopunctipennis Group was also found to be polyphyletic in the study of Harbach & Kitching (Figure 4)]. The Hyrcanus Group was paired with *An. coustani*, which corroborates previous hypotheses of a close relationship between the Hyrcanus and Coustani Groups [20,36,40,43]. Unfortunately, the analyses of Collucci & Sallum are biased by the selection of outgroup taxa whose interrelationships with the ingroup taxa were unresolved in previous studies. Thus, the results of their study cast doubt on their assertion that subgenus *Anopheles* is monophyletic. Based on the relationships recovered by Harbach & Kitching, subgenus *Anopheles* would be monophyletic if subgenus *Lophopodomyia* were to be reduced to the status of a species Group of the Anopheles Series (Figure 4). The Anopheles Series is a morphologically diverse assemblage of species and informal taxonomic groups, a number of which at one time or another were deemed to merit recognition as subgenera [20]. Sallum et al. [40] also found the Anopheles Series to be polyphyletic, but with its members interspersed in a complexity of inter-group relationships rather than arrayed in a pectinate sequence (Figure 4).

All phylogenetic studies conducted to date have demonstrated the monophyly of subgenera *Cellia* [36,38‒41], *Kerteszia* [36,38‒41,44] and *Nyssorhynchus* [36,38‒41], and the sister pairing of *Kerteszia* and *Nyssorhynchus*[36,40,41]. The sister relationship of *Cellia* and the two New World subgenera is not inconsistent with the molecular analyses of Sallum et al. [41] if *Lophopodo‐ myia* + *Bironella* is excluded from the clade that contains *Kerteszia* + *Nyssorhynchus*, but it differs markedly from the results of their earlier study based on morphology and a larger number of taxa [40], which placed *Kerteszia* + *Nyssorhynchus*, along with *An. implexus* (Christya Series), in a sister-group relationship with *Cellia* + a clade comprised of *Bironella*, *Lophopodomyia*, *Kerteszia* and *Nyssorhynchus*. *Anopheles implexus* (Christya Series) is sister to the terminal clade formed by *Kerteszia*, *Nyssorhynchus* and *Cellia* in Figure 4.

#### **4. Distribution and phylogeography of** *Anopheles*

Interpreting the current distributions of anophelines in an evolutionary context is problematic. The supercontinent of Pangaea existed in the Late Paleozoic and Early Mesozoic Eras from about 300‒200 Mya and gradually separated 200–145 Mya into the two supercontinents of Laurasia and Gondwana [45]. As noted above, evidence from DNA sequence data and fossil calibration points [34] indicates that ancestral anophelines diverged from ancestral culicines about 217 Mya (230‒192 Mya), before the complete splitting of Pangaea. If this was the case, then the separation of *Anopheles* and *Bironella* about 54 Mya (75.8‒37.1 Mya, end of the Cretaceous to near the end of the Eocene Epoch of the Cenozoic) [34] must have occurred after the separation of Gondwana into multiple continents, i.e. Africa, South America, India, Antarctica and Australia, in the Cretaceous. Atlantica (the land mass that comprised presentday South America and Africa) separated from eastern Gondwana (the land mass that comprised Antarctica, India and Australia) 150‒140 Mya. South America started to separate from Africa in a south-to-north direction during the Middle Cretaceous (about 125‒115 Mya) [46]. At the same time, Madagascar and India began to separate from Antarctica, and separated from each other 100‒90 Mya during the Cenomanian and Turonian Stages of the Late Creta‐ ceous. India continued to move northward and collided with Eurasia about 35 Mya. Laurasia split to give rise to North America/Greenland and Eurasia about 60‒55 Mya. Africa began to move northeastward toward Europe and South America moved northward to separate from Antarctica. North and South America were joined by the Isthmus of Panama during the Pliocene, approximately 3.7‒3.0 Mya.

Hyrcanus, Plumbeus, Umbrosus Groups) were recovered as monophyletic assemblages with moderate to strong support [the Pseudopunctipennis Group was also found to be polyphyletic in the study of Harbach & Kitching (Figure 4)]. The Hyrcanus Group was paired with *An. coustani*, which corroborates previous hypotheses of a close relationship between the Hyrcanus and Coustani Groups [20,36,40,43]. Unfortunately, the analyses of Collucci & Sallum are biased by the selection of outgroup taxa whose interrelationships with the ingroup taxa were unresolved in previous studies. Thus, the results of their study cast doubt on their assertion that subgenus *Anopheles* is monophyletic. Based on the relationships recovered by Harbach & Kitching, subgenus *Anopheles* would be monophyletic if subgenus *Lophopodomyia* were to be reduced to the status of a species Group of the Anopheles Series (Figure 4). The Anopheles Series is a morphologically diverse assemblage of species and informal taxonomic groups, a number of which at one time or another were deemed to merit recognition as subgenera [20]. Sallum et al. [40] also found the Anopheles Series to be polyphyletic, but with its members interspersed in a complexity of inter-group relationships rather than arrayed in a pectinate

All phylogenetic studies conducted to date have demonstrated the monophyly of subgenera *Cellia* [36,38‒41], *Kerteszia* [36,38‒41,44] and *Nyssorhynchus* [36,38‒41], and the sister pairing of *Kerteszia* and *Nyssorhynchus*[36,40,41]. The sister relationship of *Cellia* and the two New World subgenera is not inconsistent with the molecular analyses of Sallum et al. [41] if *Lophopodo‐ myia* + *Bironella* is excluded from the clade that contains *Kerteszia* + *Nyssorhynchus*, but it differs markedly from the results of their earlier study based on morphology and a larger number of taxa [40], which placed *Kerteszia* + *Nyssorhynchus*, along with *An. implexus* (Christya Series), in a sister-group relationship with *Cellia* + a clade comprised of *Bironella*, *Lophopodomyia*, *Kerteszia* and *Nyssorhynchus*. *Anopheles implexus* (Christya Series) is sister to the terminal clade formed

Interpreting the current distributions of anophelines in an evolutionary context is problematic. The supercontinent of Pangaea existed in the Late Paleozoic and Early Mesozoic Eras from about 300‒200 Mya and gradually separated 200–145 Mya into the two supercontinents of Laurasia and Gondwana [45]. As noted above, evidence from DNA sequence data and fossil calibration points [34] indicates that ancestral anophelines diverged from ancestral culicines about 217 Mya (230‒192 Mya), before the complete splitting of Pangaea. If this was the case, then the separation of *Anopheles* and *Bironella* about 54 Mya (75.8‒37.1 Mya, end of the Cretaceous to near the end of the Eocene Epoch of the Cenozoic) [34] must have occurred after the separation of Gondwana into multiple continents, i.e. Africa, South America, India, Antarctica and Australia, in the Cretaceous. Atlantica (the land mass that comprised presentday South America and Africa) separated from eastern Gondwana (the land mass that comprised Antarctica, India and Australia) 150‒140 Mya. South America started to separate from Africa in a south-to-north direction during the Middle Cretaceous (about 125‒115 Mya) [46]. At the same time, Madagascar and India began to separate from Antarctica, and separated

sequence (Figure 4).

by *Kerteszia*, *Nyssorhynchus* and *Cellia* in Figure 4.

10 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

**4. Distribution and phylogeography of** *Anopheles*

**Figure 4.** Phylogeny of subfamily Anophelinae, modified from Harbach & Kitching [36], indicating relationships within subgenus *Anopheles*. Filled circles indicate Bremer support values greater than 0.8.

Belkin [47] hypothesized that anophelines initially differentiated in the American Mediterra‐ nean Region. In concert with this postulate, Harbach & Kitching [36] suggested a possible New World origin of subfamily Anophelinae based on the basal placement of *Chagasia* relative to *Anopheles* + *Bironella* in their phylogeny of mosquito genera. Based on a phylogeny of 16 anopheline species inferred from sequences of two protein-coding nuclear genes and the Neotropical distribution of *Chagasia* and four of the seven subgenera of *Anopheles*, Krzywinski et al. [39] agreed with the hypothesis that South America was the center of origin of Anophe‐ linae. However, as will be seen below, more recent studies suggest a different scenario for the evolution of the extant groups of the subfamily. This scenario closely reflects Christophers [48] insightful observations:

Subgenus *Anopheles* appears to be the oldest of the predominant subgenera, not only on [morphological grounds], but by reason of its worldwide distribution and the greater diversity and distinctness of its forms; almost every species of the subgenus appears to be as distinctive as are the species groups of subgenus *Myzomyia* [=*Cellia*], if not more so.

*Nyssorhynchus* appears to be a Neotropical development from some pre-*Anopheles* form, whilst the group *Arribalzagia* appears to be a highly specialized development of subgenus *Anopheles*.

*Myzomyia* shows every evidence of being a new and actively disseminating branch, as is suggested by its complete ab‐ sence from the New World. Had it been once disseminated throughout North America it is unlikely that it would have been eliminated from the whole continent so completely as to leave not a single species in this area, though there is no actual proof that this did not occur. The apparent affinity between the group *Neomyzomyia* and subgenus *Nyssorhyn‐ chus* suggests an intermediate ancestor, though not necessarily one in the south, *i. e.*, such affinity does not prove or suggest a land-connection between Australia and South America, as the common ancestor may have been derived from the north and later eliminated. [next paragraph omitted]

The date of isolation of South America, judging by the history of mammals, would be from the middle of the Eocene, when connections between North and South America were severed, until the end of the Pliocene (*Zittel*). The anophe‐ line fauna, therefore, arose from elements which pre-dated this period, and there were already subgenus *Anopheles*-like forms, as well as some earlier type from which *Nyssorhynchus* arose.

At some unknown period a similar special development took place, resulting in an early form (*Neomyzomyia*) of subge‐ nus *Myzomyia*. This form appears to have once been distributed throughout the Oriental, Ethiopian [i.e. Afrotropical], and Australian Regions, and to have later undergone some regression, eventually remaining in greatest strength in the Australian Region.

Edwards, in reviewing the fossil remains of mosquitoes, notes that probably all the main divisions of the family [Culi‐ cidae] existed in Mid-Tertiary much as they do today, and with almost identical characters, and considers that, though no fossil *Anopheles* have been found, there can be no doubt from its morphology that this is also an old genus, probably older than any culicine form.

Based on the relationships shown in Figure 4, distributions of the principal group taxa (Appendix 6) and the geological dates listed above, it would appear that the ancestral lineage of *Anopheles* existed before the breakup of Pangaea and subsequently diversified into the modern subgenera and species after the separation of the continents. This would explain the cosmopolitan distribution and greater diversity of subgenus *Anopheles*, but not the earlier divergence of genus *Chagasia* and subgenus *Stethomyia*, which are confined to the Neotropical Region, the Oriental subgenus *Baimaia* and the Australasian genus *Bironella* (Figure 4). *Chagasia* possess several features that characterize species of subfamily Culicinae, including the strongly arched mesonotum, trilobed scutellum (Figure 6) and setae on the postpronotum. Based on these shared features, *Chagasia* has been considered an ancient group showing affinities with non-anophelines and phylogenetic analyses of morphological data and DNA sequences of various ribosomal, mitochondrial and nuclear genes strongly support its placement in an ancestral relationship to all other anophelines [33,35‒41]. From the foregoing, however, it is inferred here that *Chagasia*, with only seven species, is a relic of a once more widely distributed taxon that is now confined to residual areas of South and Central America. It is also possible, although less likely, that *Chagasia*, as suggested by the late John N. Belkin for other mosquitoes [47], may have originated through hybridization between early anopheline and culicine forms.

Subgenus *Anopheles* appears to be the oldest of the predominant subgenera, not only on [morphological grounds], but by reason of its worldwide distribution and the greater diversity and distinctness of its forms; almost every species of the subgenus appears to be as distinctive as are the species groups of subgenus *Myzomyia* [=*Cellia*], if not more so. *Nyssorhynchus* appears to be a Neotropical development from some pre-*Anopheles* form, whilst the group *Arribalzagia*

*Myzomyia* shows every evidence of being a new and actively disseminating branch, as is suggested by its complete ab‐ sence from the New World. Had it been once disseminated throughout North America it is unlikely that it would have been eliminated from the whole continent so completely as to leave not a single species in this area, though there is no actual proof that this did not occur. The apparent affinity between the group *Neomyzomyia* and subgenus *Nyssorhyn‐ chus* suggests an intermediate ancestor, though not necessarily one in the south, *i. e.*, such affinity does not prove or suggest a land-connection between Australia and South America, as the common ancestor may have been derived

The date of isolation of South America, judging by the history of mammals, would be from the middle of the Eocene, when connections between North and South America were severed, until the end of the Pliocene (*Zittel*). The anophe‐ line fauna, therefore, arose from elements which pre-dated this period, and there were already subgenus *Anopheles*-like

At some unknown period a similar special development took place, resulting in an early form (*Neomyzomyia*) of subge‐ nus *Myzomyia*. This form appears to have once been distributed throughout the Oriental, Ethiopian [i.e. Afrotropical], and Australian Regions, and to have later undergone some regression, eventually remaining in greatest strength in the

Edwards, in reviewing the fossil remains of mosquitoes, notes that probably all the main divisions of the family [Culi‐ cidae] existed in Mid-Tertiary much as they do today, and with almost identical characters, and considers that, though no fossil *Anopheles* have been found, there can be no doubt from its morphology that this is also an old genus, probably

Based on the relationships shown in Figure 4, distributions of the principal group taxa (Appendix 6) and the geological dates listed above, it would appear that the ancestral lineage of *Anopheles* existed before the breakup of Pangaea and subsequently diversified into the modern subgenera and species after the separation of the continents. This would explain the cosmopolitan distribution and greater diversity of subgenus *Anopheles*, but not the earlier divergence of genus *Chagasia* and subgenus *Stethomyia*, which are confined to the Neotropical Region, the Oriental subgenus *Baimaia* and the Australasian genus *Bironella* (Figure 4). *Chagasia* possess several features that characterize species of subfamily Culicinae, including the strongly arched mesonotum, trilobed scutellum (Figure 6) and setae on the postpronotum. Based on these shared features, *Chagasia* has been considered an ancient group showing affinities with non-anophelines and phylogenetic analyses of morphological data and DNA sequences of various ribosomal, mitochondrial and nuclear genes strongly support its placement in an ancestral relationship to all other anophelines [33,35‒41]. From the foregoing, however, it is inferred here that *Chagasia*, with only seven species, is a relic of a once more widely distributed taxon that is now confined to residual areas of South and Central America. It is also possible, although less likely, that *Chagasia*, as suggested by the late John N. Belkin for other mosquitoes [47], may have originated through hybridization between early anopheline and culicine forms.

appears to be a highly specialized development of subgenus *Anopheles*.

12 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

from the north and later eliminated. [next paragraph omitted]

forms, as well as some earlier type from which *Nyssorhynchus* arose.

Australian Region.

older than any culicine form.

**Figure 5.** Phylogeny of subgenera *Cellia*, *Kerteszia* and *Nyssorhynchus*, modified from Harbach & Kitching [36], indicat‐ ing relationships within subgenera *Cellia* and *Nyssorhynchus*. Filled circles indicate Bremer support values greater than 0.8.

Similarly, *Bironella* (as suggested by Christophers [48]), *Baimaia* and *Stethomyia*, with few species and restricted distributions, are also the remnants of once much more widely distrib‐ uted forms. The isolation of ancestral members of subgenus *Anopheles* in South America also explains the uniqueness of the extant Neotropical fauna of the subgenus, especially the welldifferentiated Arribalzagia Series. In accordance with this hypothesis, the following groups are also probably residual elements of once more widely distributed ancestral forms of subgenus *Anopheles*: the Afrotropical Christya Series (two species), the Australasian Atratipes (two species) and Stigmaticus (six species) Groups, the Oriental Alongensis (two species) and Culiciformis (three species) Groups, the Oriental Lophoscelomyia Series (five species) and the Neotropical Cycloleppteron Series (two species). It is noteworthy that the extant members of the relict groups are not vectors of human malarial parasites.

As noted previously, subgenus *Anopheles* has an almost world-wide distribution. Species are found at elevations from coastal areas to mountainous terrain in temperate, subtropical and tropical areas, but are absent from the majority of the Pacific Islands, including the large ones of New Zealand, Fiji and New Caledonia. The sole species of subgenus *Baimaia* has been found only in forested hilly and mountainous areas between 14° and 17° north on either side of the Thai-Myanmar border and at a location near the Thai-Laos border in Thailand, and is probably also a relict taxon that has retained generalized ancestral features of the male genitalia [36]. Most species of subgenus *Cellia* have distributions in the Afrotropical, Australasian and Oriental Regions, but some species occur in southern areas of the Palaearctic. Species of *Cellia* are conspicuously absent from the majority of the islands of the Pacific, including New Zealand, Fiji and New Caledonia. Species of subgenus *Kerteszia* are found in the Neotropical Region, from Veracruz State in Mexico through Central America and Atlantic South America, along the Andes and along the coast, to the States of Misiones in Argentina and Rio Grande do Sul in Brazil, and also occur south along the Pacific Coast of South America to the State of El Oro, Ecuador. The subgenus is absent from all islands of the West Indies except Trinidad, and from most of the vast expanse of the Amazon basin in South America [49]. Species of subgenus *Lophopodomyia* are known to occur in areas of Panama and northern South America (Brazil, Colombia, Ecuador, French Guiana and Venezuela). Species of subgenus *Nyssorhyn‐ chus* are restricted to the Neotropical Region, except for *An. albimanus*, which extends into the Nearctic Region (northern Mexico and along the Rio Grande River in Texas). Finally, species of subgenus *Stethomyia* principally occur in southern Central America (Costa Rica and Panama) and northern South America (Brazil, Colombia, French Guiana, Guyana, Suriname and Venezuela), but one or two species are known to occur on the islands of Trinidad and Tobago and as far south as Peru and Bolivia.

**Figure 6.** Two forms of the mosquito scutellum (Stm): A, trilobed scutellum of *Chagasia* and species of subfamily Culi‐ cinae; B, evenly rounded scutellum of *Anopheles*, with few exceptions. Original images from Harbach & Kitching [18].

Subgenera *Kerteszia*, *Lophopodomyia*, *Nyssorhynchus* and *Stethomyia*, and the Arribalzagia and Cycloleppteron Series of subgenus *Anopheles* are special to the Neotropical Region, where they probably originated following the separation of South America and Africa. The derived position of subgenera *Cellia* and *Kerteszia* + *Nyssorhynchus* relative to subgenus *Anopheles* (Figure 4) supports the hypothesis that the stem lineage of these subgenera originated in Gondwana and diverged following the separation of Atlantica to give rise to *Cellia* in Africa and *Kerteszia* and *Nyssorhynchus* in South America. It is interesting to note that *Lophopodo‐* *myia* and the Pseudopunctipennis Group are sister taxa in Figure 4, which is plausible in view of the hypothesized evolution of these groups from Neotropical ancestors. The Pseudopunc‐ tipennis Group is nearly restricted to the Neotropics, except for *An. franciscanus* and a minor extension of *An. pseudopunctipennis* into the Nearctic Region, which undoubtedly occurred relatively recently, after the land bridge formed to connect North and South America 3.7‒3.0 Mya. Except for these two species, all *Anopheles* species in the Nearctic Region are members of the Anopheles Series of subgenus *Anopheles*. Half of the species of the Holarctic Maculipennis Group (24 species) occur in the Nearctic Region and the other half occur in the Palaearctic. This indicates that the Maculipennis Group must have evolved in the Northern Hemisphere prior to the separation of North America and Eurasia during the Paleocene and Eocene Epochs (60‒ 55 Mya). The Plumbeus Group includes species in the Nearctic (2), Neotropical (4) and Palaearctic (3) Regions. Its position in the cladogram shown in Figure 4 is based on *An. judithae*, a Nearctic species. This group may be what paleontologists call a "stem group" [50], a paraphyletic or polyphyletic assemblage of species that share features of extinct taxa. The spotted distribution of these "living fossil" species suggests that their extinct relatives, ancestral forms of the Anopheles Series, existed before the separation of Pangaea. This bodes well with Christophers & Barraud's 1931 hypothesis [51] that the eggs of species of the Plumbeus Group are primitive compared to other species of subgenus *Anopheles*.

Thai-Myanmar border and at a location near the Thai-Laos border in Thailand, and is probably also a relict taxon that has retained generalized ancestral features of the male genitalia [36]. Most species of subgenus *Cellia* have distributions in the Afrotropical, Australasian and Oriental Regions, but some species occur in southern areas of the Palaearctic. Species of *Cellia* are conspicuously absent from the majority of the islands of the Pacific, including New Zealand, Fiji and New Caledonia. Species of subgenus *Kerteszia* are found in the Neotropical Region, from Veracruz State in Mexico through Central America and Atlantic South America, along the Andes and along the coast, to the States of Misiones in Argentina and Rio Grande do Sul in Brazil, and also occur south along the Pacific Coast of South America to the State of El Oro, Ecuador. The subgenus is absent from all islands of the West Indies except Trinidad, and from most of the vast expanse of the Amazon basin in South America [49]. Species of subgenus *Lophopodomyia* are known to occur in areas of Panama and northern South America (Brazil, Colombia, Ecuador, French Guiana and Venezuela). Species of subgenus *Nyssorhyn‐ chus* are restricted to the Neotropical Region, except for *An. albimanus*, which extends into the Nearctic Region (northern Mexico and along the Rio Grande River in Texas). Finally, species of subgenus *Stethomyia* principally occur in southern Central America (Costa Rica and Panama) and northern South America (Brazil, Colombia, French Guiana, Guyana, Suriname and Venezuela), but one or two species are known to occur on the islands of Trinidad and Tobago

**Figure 6.** Two forms of the mosquito scutellum (Stm): A, trilobed scutellum of *Chagasia* and species of subfamily Culi‐ cinae; B, evenly rounded scutellum of *Anopheles*, with few exceptions. Original images from Harbach & Kitching [18].

Subgenera *Kerteszia*, *Lophopodomyia*, *Nyssorhynchus* and *Stethomyia*, and the Arribalzagia and Cycloleppteron Series of subgenus *Anopheles* are special to the Neotropical Region, where they probably originated following the separation of South America and Africa. The derived position of subgenera *Cellia* and *Kerteszia* + *Nyssorhynchus* relative to subgenus *Anopheles* (Figure 4) supports the hypothesis that the stem lineage of these subgenera originated in Gondwana and diverged following the separation of Atlantica to give rise to *Cellia* in Africa and *Kerteszia* and *Nyssorhynchus* in South America. It is interesting to note that *Lophopodo‐*

and as far south as Peru and Bolivia.

14 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Species in subgenus *Cellia* are confined to the Eastern Hemisphere, with members in the Afrotropical, Australasian, Oriental and Palaearctic regions (Figure 5, Appendix 6). The Afrotropical Region is characterized by a large number of species of subgenus *Cellia* and relatively few species of subgenus *Anopheles*. The Myzomyia Series is especially dominant, but species of the Neocellia, Neomyzomyia and Pyretophorus Series also occur in the region. The Myzomyia, Neocellia and Pyretophorus Series are represented in the Afrotropical and Oriental Regions, but no species, species groups or subgroups of these series (with the exception of the Minimus Subgroup) are common to both regions (see Appendix 6). The Myzomyia Series is a dominant group in Africa, where *An. funestus* is a principal malaria vector [52,53]. Related species of the Funestus Group, including *An. minimus* and other members of the Minimus Subgroup, are major vectors of malarial parasites in southern Asia [52,54]. Evidence from phylogenetic analyses of mitochondrial DNA (ITS2 and D3 sequences) indicates that the Funestus Group originated in the Afrotropical Region [55]. The Neocellia Series also includes several important malaria vectors in southern Asia, notably *An. stephensi* and members of the Maculatus Group [52,54]. The Pyretophorus Series includes the formidable malaria vectors of the Gambiae Complex in Africa and important vectors of the Sundaicus and Subpictus Complexes in Southeast Asia [53,54]. The morphology-based phylogeny of Anthony et al. [56] indicates that the Pyretophorus Series originated in Africa and suggests that the capacity to vector malarial parasites is an ancestral condition subsequently lost independently in several lineages.

The anopheline fauna of the Australasian Region also shows evidence of isolation, but not to the degree indicated by the Neotropical fauna. The isolation appears to be more recent, corresponding to the separation of Australia from Antarctica between 37.0‒33.5 Mya. The region includes a preponderance of species of the Neomyzomyia Series of subgenus *Cellia*, which may signal a relatively recent arrival from the Oriental Region, with some diversifica‐ tion. Members of the Neomyzomyia Series are the only *Anopheles* in the South Pacific [47]. Species groups of the series are confined to the Afrotropical (Ardensis, Mascarensis, Pauliani, Ranci, Rhodesiensis and Smithii Groups), Australasian (Punctulatus Group, Lungae Complex and unassigned species) or Oriental Region (Kochi, Leucosphyrus and Tessellatus Groups) (Appendix 6). The Neomyzomyia Series has been regarded as the most primitive series of subgenus *Cellia* based on egg morphology and the reduced or non-existent cibarial armature of females [57‒59], and is thought to have originated in Africa and subsequently disperse eastward to the Oriental and Australasian Regions [52,59]. None of the African species of the Neomyzomyia Series, except for *An. nili*, are major vectors of malaria. In comparison, most species of the Oriental Leucosphyrus and Australasian Punctulatus Groups of the Neomyzo‐ myia Series are important vectors of both primate and human malarial parasites. The Cellia and Paramyzomyia Series of subgenus *Cellia* are restricted to the Afrotropical Region, except for *An. pharoensis* (Cellia Series) and *An. multicolor* (Paramyzomyia Series) which occur in adjacent arid areas of the Palaearctic (Sahara and Middle East). It seems reasonable to hy‐ pothesize that those series that are presently represented by groups in the Afrotropical, Australasian and Oriental Regions arose before eastern Gondwana (Antarctica, India and Australia) fragmented. The Mascarensis, Pauliani and Ranci Groups are confined to Mada‐ gascar, which supports the hypothesis that the ancestral forms of at least these groups of the Neomyzomyia Series existed before Madagascar separated from India 100‒90 Mya.

Human malaria probably evolved in Africa along with its mosquito hosts and other primates. Modern humans arose in Africa about 200,000 years ago and dispersed into Eurasia [60], reaching Australia about 40,000 years ago. Migration into the New World occurred about 15‒ 20 millennia ago, and most of the Pacific Islands were colonized by four thousand years ago. The point here is that the rise and dispersal of modern humans occurred long after the formation of the continents and the evolution of the major groups of *Anopheles*. Consequently, it seems reasonable to assume that human malarial parasites accompanied humans during their migration out of Africa and were passed on to species of *Anopheles* in other regions that had the ecological, physiological and behavioural attributes required to propagate infections and maintain transmission. These taxa were surely already adapted to feeding on primates, including the ancestors of *Homo sapiens*, and were capable of developing and transmitting the *Plasmodium* species specific to those hosts.

Comprehensive information on the dominant malaria vectors of the world, most of which are presumably recently evolved members of sibling species complexes (Appendix 3), is sum‐ marized in a series of publications (and a chapter of this book) by M. Sinka and a team of regional experts and technical advisors ‒ the Americas [61], Africa, Europe and the Middle East [53], the Asia-Pacific Region [54] ‒ that culminated in a thorough review of the principal malaria vector taxa of the world [62]. At present, 96 formally named species of *Anopheles* are members of 26 sibling species complexes (Appendix 4). Twenty of these nominal species actually consist of more than one species, which all together comprise a total of 67 species. Excluding the name-bearing type species, the 58 species, plus five other unnamed species that are not members of species complexes, a total of 72 species, have yet to be given formal Latin names (Appendix 4).

## **5. Conclusion**

which may signal a relatively recent arrival from the Oriental Region, with some diversifica‐ tion. Members of the Neomyzomyia Series are the only *Anopheles* in the South Pacific [47]. Species groups of the series are confined to the Afrotropical (Ardensis, Mascarensis, Pauliani, Ranci, Rhodesiensis and Smithii Groups), Australasian (Punctulatus Group, Lungae Complex and unassigned species) or Oriental Region (Kochi, Leucosphyrus and Tessellatus Groups) (Appendix 6). The Neomyzomyia Series has been regarded as the most primitive series of subgenus *Cellia* based on egg morphology and the reduced or non-existent cibarial armature of females [57‒59], and is thought to have originated in Africa and subsequently disperse eastward to the Oriental and Australasian Regions [52,59]. None of the African species of the Neomyzomyia Series, except for *An. nili*, are major vectors of malaria. In comparison, most species of the Oriental Leucosphyrus and Australasian Punctulatus Groups of the Neomyzo‐ myia Series are important vectors of both primate and human malarial parasites. The Cellia and Paramyzomyia Series of subgenus *Cellia* are restricted to the Afrotropical Region, except for *An. pharoensis* (Cellia Series) and *An. multicolor* (Paramyzomyia Series) which occur in adjacent arid areas of the Palaearctic (Sahara and Middle East). It seems reasonable to hy‐ pothesize that those series that are presently represented by groups in the Afrotropical, Australasian and Oriental Regions arose before eastern Gondwana (Antarctica, India and Australia) fragmented. The Mascarensis, Pauliani and Ranci Groups are confined to Mada‐ gascar, which supports the hypothesis that the ancestral forms of at least these groups of the

Neomyzomyia Series existed before Madagascar separated from India 100‒90 Mya.

*Plasmodium* species specific to those hosts.

16 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Human malaria probably evolved in Africa along with its mosquito hosts and other primates. Modern humans arose in Africa about 200,000 years ago and dispersed into Eurasia [60], reaching Australia about 40,000 years ago. Migration into the New World occurred about 15‒ 20 millennia ago, and most of the Pacific Islands were colonized by four thousand years ago. The point here is that the rise and dispersal of modern humans occurred long after the formation of the continents and the evolution of the major groups of *Anopheles*. Consequently, it seems reasonable to assume that human malarial parasites accompanied humans during their migration out of Africa and were passed on to species of *Anopheles* in other regions that had the ecological, physiological and behavioural attributes required to propagate infections and maintain transmission. These taxa were surely already adapted to feeding on primates, including the ancestors of *Homo sapiens*, and were capable of developing and transmitting the

Comprehensive information on the dominant malaria vectors of the world, most of which are presumably recently evolved members of sibling species complexes (Appendix 3), is sum‐ marized in a series of publications (and a chapter of this book) by M. Sinka and a team of regional experts and technical advisors ‒ the Americas [61], Africa, Europe and the Middle East [53], the Asia-Pacific Region [54] ‒ that culminated in a thorough review of the principal malaria vector taxa of the world [62]. At present, 96 formally named species of *Anopheles* are members of 26 sibling species complexes (Appendix 4). Twenty of these nominal species actually consist of more than one species, which all together comprise a total of 67 species. Excluding the name-bearing type species, the 58 species, plus five other unnamed species that

A more robust phylogeny of *Anopheles* mosquitoes than is currently available may be of use in the fight against malaria. Foley et al. [37] suggested that it may help "by elucidating descent relationships of genes for refractoriness, insecticide resistance, and genetically determined ecological and behavioral traits important to malaria transmission." Interrupting the life cycle of malarial parasites by genetically manipulating vector receptiveness to infection is a potential approach to malaria control. A natural classification of *Anopheles* predictive of biological and ecological traits could facilitate the manipulation of vector genomes by informing the dynamics of introduced genes. Obviously, co-evolutionary studies of parasites and vectors require phylogenies for the mosquitoes. This must far exceed the taxon-limited (exemplar-based) studies conducted to date as they do not provide a basis for gaining insights into interspecific and co-evolutionary relationships of vectors and parasites.

It seems fitting to end here with a comment concerning interspecific hybridization, which was mentioned above in relation to genus *Chagasia* in the Neotropical Region. Although anopheline species occur in sympatry in most ecosystems, hybridization has only been detected at very low levels between certain members of species complexes in subgenus *Cellia*, e.g. *An. gam‐ biae* with both *An. arabiensis* and *An. bwambae* in Africa [63,64], *An. dirus* and *An. baimaii* in Thailand [65] and *An. minimus* and *An. harrisoni* in Vietnam [66]. However, as advocated by Belkin [47], hybridization could provide sufficient genetic variation to permit adaptation to new habitats. Hybridization may occur regularly between some species, particularly widely distributed species that are morphologically similar. It could have played a role in the speciation and evolution of *Anopheles* mosquitoes and the pathogens they transmit.


## **Appendix 1 — The internal classification of genus** *Anopheles*



**Subgenus Section Series Group Subgroup Complex Author**

18 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Pseudopunctipen‐

Laticorn [20]

nis

Culiciformis [20] Lindesayi [20]

Maculipennis [20]

Plumbeus [20]

Punctipennis [20]

Stigmaticus [20]

Asiaticus [23]

Albotaeniatus [20] Bancroftii [20] Barbirostris [20]

Coustani [20] Hyrcanus [77]

Asiaticus [73] Interruptus [73]

Barbirostris [23]

Vanus [23]

Lesteri [78] Nigerrimus [78]

Barbirostris [76]

Cycloleppteron [12] Lophoscelomyia [12]

Arribalzagia [74] Christya [75] Myzorhynchus [12]

Maculipennis [71] Quadrimaculatus [71] Freeborni [71]

Gigas [70] Lindesayi [70]

Crucians [72]

[20]



**Subgenus Section Series Group Subgroup Complex Author**

20 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Longirostris [90] Lungae [47]

Nili [22]

Dirus [94] Leucosphyrus [92]

Farauti [96]

Gambiae [97]

Sundaicus [98]

Subpictus [99]

Cruzii [101]

Ardensis [22]

Kochi [73] Leucosphyrus [91]

Mascarensis [26] Pauliani [21] Punctulatus [95]

Ranci [21]

Rhodesiensis [22] Smithii [22] Tessellatus [73]

Cinereus [22] Listeri [22]

Ludlowae [73]

Subpictus [73]

Paramyzomyia [51]

Pyretophorus [12]

*Kerteszia* [100]

Hackeri [92] Leucosphyrus [93]

Riparis [93]

Ranci [21] Roubaudi [21]

> Alphabetical list of formally named species of *Anopheles* and their position in the classification of the genus. For species Complexes, see Appendices 3 and 4; for authorship of species, visit http://mosquito-taxonomic-inventory.info/valid-species-list.




**Species Subgenus Section Series Group Subgroup**

*aconitus Cellia* Myzomyia Funestus Aconitus *ahomi Anopheles* Laticorn Myzorhynchus Barbirostris Vanus

*albertoi Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Strodei

*anomalophyllus Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*aquasalis Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*artemievi Anopheles* Angusticorn Anopheles Maculipennis Maculipennis

*acaci Anopheles* Angusticorn Anopheles Aitkenii

*aitkenii Anopheles* Angusticorn Anopheles Aitkenii

*albitarsis Nyssorhynchus* Argyritarsis Albitarsis Albitarsis *albotaeniatus Anopheles* Laticorn Myzorhynchus Albotaeniatus

*alongensis Anopheles* Angusticorn Anopheles Alongensis

*annandalei Anopheles* Angusticorn Lophoscelomyia Asiaticus *annularis Cellia* Neocellia Annularis

*ainshamsi Cellia* Neocellia

*albimanus Nyssorhynchus* Albimanus Albimanus

*algeriensis Anopheles* Angusticorn Anopheles

*amictus Cellia* Neomyzomyia *anchietai Anopheles* Laticorn Arribalzagia

*annulatus Cellia* Neomyzomyia *annulipalpis Anopheles* Angusticorn Cycloleppteron *annulipes Cellia* Neomyzomyia

*apicimacula Anopheles* Laticorn Arribalzagia *apoci Cellia* Myzomyia

*arabiensis Cellia* Pyretophorus

*argenteolobatus Cellia* Cellia

*arboricola Anopheles* Angusticorn Anopheles Plumbeus *ardensis Cellia* Neomyzomyia Ardensis

*argyritarsis Nyssorhynchus* Argyritarsis Argyritarsis Argyritarsis *argyropus Anopheles* Laticorn Myzorhynchus Hyrcanus

*antunesi Nyssorhynchus* Myzorhynchella

*acanthotorynus Stethomyia*

22 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors



**Species Subgenus Section Series Group Subgroup**

*brucei Cellia* Myzomyia Funestus Rivulorum

*campestris Anopheles* Laticorn Myzorhynchus Barbirostris Barbirostris

*berghei Cellia* Myzomyia Marshallii

*borneensis Anopheles* Angusticorn Anopheles Aitkenii *bradleyi Anopheles* Angusticorn Anopheles Punctipennis *braziliensis Nyssorhynchus* Argyritarsis Albitarsis Braziliensis *brevipalpis Anopheles* Laticorn Myzorhynchus Umbrosus *brevirostris Anopheles* Laticorn Myzorhynchus Umbrosus *brohieri Cellia* Myzomyia Marshallii

*buxtoni Cellia* Neomyzomyia Ardensis

*caliginosus Anopheles* Laticorn Myzorhynchus Coustani *cameroni Cellia* Neomyzomyia Rhodesiensis

*canorii Stethomyia* Neomyzomyia Smithii *carnevalei Cellia* Neomyzomyia Ardensis

*carteri Cellia* Myzomyia Demeilloni

*chodukini Anopheles* Laticorn Myzorhynchus Hyrcanus

*cinctus Cellia* Neomyzomyia Ardensis *cinereus Cellia* Paramyzomyia Cinereus

*clowi Cellia* Neomyzomyia Punctulatus

*christyi Cellia* Pyretophorus

*claviger Anopheles* Angusticorn Anopheles

*chiriquiensis Anopheles* Angusticorn Anopheles Pseudopunctipennis

*bwambae Cellia* Pyretophorus *calderoni Anopheles* Laticorn Arribalzagia

*bervoetsi Cellia* Myzomyia

24 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

*brumpti Cellia* Cellia *brunnipes Cellia* Myzomyia *bulkleyi Anopheles* Angusticorn Lophoscelomyia *bustamentei Anopheles* Laticorn Arribalzagia

*boliviensis Kerteszia*

*caroni Cellia*



**Species Subgenus Section Series Group Subgroup** *donaldi Anopheles* Laticorn Myzorhynchus Barbirostris Barbirostris *dravidicus Cellia* Neocellia Maculatus Maculatus

*dunhami Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*earlei Anopheles* Angusticorn Anopheles Maculipennis Freeborni

*elegans Cellia* Neomyzomyia Leucosphyrus Leucosphyrus

*evansae Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*filipinae Cellia* Myzomyia Funestus Aconitus

*flavirostris Cellia* Myzomyia Funestus Minimus

*fluviatilis Cellia* Myzomyia Funestus Minimus

*franciscoi Anopheles* Laticorn Myzorhynchus Barbirostris Barbirostris

*eiseni Anopheles* Angusticorn Anopheles Pseudopunctipennis *ejercitoi Anopheles* Laticorn Myzorhynchus Albotaeniatus

*dthali Cellia* Myzomyia *dualaensis Cellia* Neomyzomyia

26 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

*dureni Cellia* Neomyzomyia Ardensis

*engarensis Anopheles* Laticorn Myzorhynchus Hyrcanus *eouzani Cellia* Neomyzomyia Ardensis

*erepens Cellia* Myzomyia Wellcomei

*faini Cellia* Neomyzomyia Smithii *farauti Cellia* Neomyzomyia Punctulatus *fausti Anopheles* Angusticorn Anopheles Plumbeus

*epiroticus Cellia* Pyretophorus

*erythraeus Cellia* Myzomyia *ethiopicus Cellia* Myzomyia *evandroi Anopheles* Laticorn Arribalzagia

*flavicosta Cellia* Myzomyia

*fluminensis Anopheles* Laticorn Arribalzagia

*fontinalis Cellia* Myzomyia *forattinii Anopheles* Laticorn Arribalzagia

*fragilis Anopheles* Angusticorn Anopheles Aitkenii

*franciscanus Anopheles* Angusticorn Anopheles Pseudopunctipennis



**Species Subgenus Section Series Group Subgroup** *harrisoni Cellia* Myzomyia Funestus Minimus

*hermsi Anopheles* Angusticorn Anopheles Maculipennis Freeborni

*hodgkini Anopheles* Laticorn Myzorhynchus Barbirostris Barbirostris

*ininii Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*interruptus Anopheles* Angusticorn Lophoscelomyia Asiaticus Interruptus *introlatus Cellia* Neomyzomyia Leucosphyrus Leucosphyrus *inundatus Anopheles* Angusticorn Anopheles Maculipennis Quadrimaculatus

*hectoris Anopheles* Angusticorn Anopheles Pseudopunctipennis

*heiheensis Anopheles* Laticorn Myzorhynchus Hyrcanus

*hinesorum Cellia* Neomyzomyia Punctulatus

*hughi Cellia* Myzomyia Marshallii *hunteri Anopheles* Laticorn Myzorhynchus Umbrosus *hyrcanus Anopheles* Laticorn Myzorhynchus Hyrcanus

*indefinitus Cellia* Pyretophorus Subpictus

*insulaeflorum Anopheles* Angusticorn Anopheles Aitkenii

*irenicus Cellia* Neomyzomyia Punctulatus *jamesii Cellia* Neocellia Jamesii *janconnae Nyssorhynchus* Argyritarsis Albitarsis Albitarsis *jebudensis Cellia* Neomyzomyia Smithii *jeyporiensis Cellia* Myzomyia Funestus *judithae Anopheles* Angusticorn Anopheles Plumbeus

*keniensis Cellia* Myzomyia Demeilloni *kingi Cellia* Neomyzomyia Ardensis *kleini Anopheles* Laticorn Myzorhynchus Hyrcanus

*hervyi Cellia* Neocellia *hilli Cellia* Neomyzomyia

28 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

*implexus Anopheles* Laticorn Christya *incognitus Cellia* Neomyzomyia

*intermedius Anopheles* Laticorn Arribalzagia

*karwari Cellia* Neocellia

*homunculus Kerteszia*



**Species Subgenus Section Series Group Subgroup**

*macarthuri Cellia* Neomyzomyia Leucosphyrus Riparis

*maculatus Cellia* Neocellia Maculatus Maculatus

*maculipennis Anopheles* Angusticorn Anopheles Maculipennis Maculipennis

*manalangi Anopheles* Laticorn Myzorhynchus Barbirostris Vanus *mangyanus Cellia* Myzomyia Funestus Aconitus

*martinius Anopheles* Angusticorn Anopheles Maculipennis Maculipennis

*maverlius Anopheles* Angusticorn Anopheles Maculipennis Quadrimaculatus

*melanoon Anopheles* Angusticorn Anopheles Maculipennis Maculipennis

*messeae Anopheles* Angusticorn Anopheles Maculipennis Maculipennis

*lovettae Cellia* Neomyzomyia Smithii *ludlowae Cellia* Pyretophorus Ludlowae

*machardyi Cellia* Neomyzomyia Ardensis

*maliensis Cellia* Neomyzomyia Ardensis

*marajoara Nyssorhynchus* Argyritarsis Albitarsis Albitarsis *marshallii Cellia* Myzomyia Marshallii

*mascarensis Cellia* Neomyzomyia Mascarensis

*mengalangensis Anopheles* Angusticorn Anopheles Lindesayi

*millecampsi Cellia* Neomyzomyia Ardensis *milloti Cellia* Neomyzomyia Pauliani

*lungae Cellia* Neomyzomyia

*maculipalpis Cellia* Neocellia

*maculipes Anopheles* Laticorn Arribalzagia *majidi Cellia* Myzomyia *malefactor Anopheles* Laticorn Arribalzagia

*marteri Anopheles* Angusticorn Anopheles

*mattogrossensis Anopheles* Laticorn Arribalzagia

*mediopunctatus Anopheles* Laticorn Arribalzagia

*melas Cellia* Pyretophorus

*meraukensis Cellia* Neomyzomyia *merus Cellia* Pyretophorus

*lutzii Nyssorhynchus* Myzorhynchella

30 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors



**Species Subgenus Section Series Group Subgroup**

*occidentalis Anopheles* Angusticorn Anopheles Maculipennis Freeborni

*oswaldoi Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*pampanai Cellia* Myzomyia Funestus Aconitus

*paraliae Anopheles* Laticorn Myzorhynchus Hyrcanus Lesteri

*parensis Cellia* Myzomyia Funestus Funestus

*peditaeniatus Anopheles* Laticorn Myzorhynchus Hyrcanus Lesteri

*persiensis Anopheles* Angusticorn Anopheles Maculipennis Maculipennis

*parapunctipennis Anopheles* Angusticorn Anopheles Pseudopunctipennis

*obscurus Anopheles* Laticorn Myzorhynchus

32 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

*okuensis Anopheles* Laticorn Christya

*paltrinierii Cellia* Neocellia

*parangensis Cellia* Pyretophorus

*pattoni Cellia* Neocellia

*peryassui Anopheles* Laticorn Arribalzagia *petragnani Anopheles* Angusticorn Anopheles

*pharoensis Cellia* Cellia

*pholidotus Kerteszia*

*parvus Nyssorhynchus* Myzorhynchella

*omorii Anopheles* Angusticorn Anopheles Plumbeus *oryzalimnetes Nyssorhynchus* Argyritarsis Albitarsis Albitarsis

*ovengensis Cellia* Neomyzomyia Ardensis *pallidus Cellia* Neocellia Annularis *palmatus Anopheles* Angusticorn Anopheles Aitkenii

*paludis Anopheles* Laticorn Myzorhynchus Coustani

*papuensis Anopheles* Angusticorn Anopheles Stigmaticus

*pauliani Cellia* Neomyzomyia Pauliani

*peytoni Anopheles* Angusticorn Anopheles Aitkenii

*philippinensis Cellia* Neocellia Annularis

*pictipennis Nyssorhynchus* Argyritarsis Argyritarsis Pictipennis

*perplexens Anopheles* Angusticorn Anopheles Punctipennis

*oiketorakras Lophopodomyia*



**Species Subgenus Section Series Group Subgroup** *ranci Cellia* Neomyzomyia Ranci Ranci *rangeli Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi *recens Cellia* Neomyzomyia Leucosphyrus Hackeri *reidi Anopheles* Laticorn Myzorhynchus Barbirostris Vanus

*riparis Cellia* Neomyzomyia Leucosphyrus Riparis *rivulorum Cellia* Myzomyia Funestus Rivulorum

*rondoni Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Strodei *roperi Anopheles* Laticorn Myzorhynchus Umbrosus Letifer *roubaudi Cellia* Neomyzomyia Ranci Roubaudi

*sacharovi Anopheles* Angusticorn Anopheles Maculipennis Maculipennis

*sanctielii Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*sawadwongporni Cellia* Neocellia Maculatus Sawadwongporni

*scanloni Cellia* Neomyzomyia Leucosphyrus Leucosphyrus

*separatus Anopheles* Laticorn Myzorhynchus Umbrosus Separatus

*rennellensis Cellia* Neomyzomyia Punctulatus *rhodesiensis Cellia* Neomyzomyia Rhodesiensis

*ruarinus Cellia* Neomyzomyia Rhodesiensis

*samarensis Anopheles* Laticorn Myzorhynchus Umbrosus

*saperoi Anopheles* Laticorn Myzorhynchus Albotaeniatus

*sawyeri Nyssorhynchus* Argyritarsis Argyritarsis Argyritarsis

*schueffneri Cellia* Neocellia Annularis

*seretsei Cellia* Paramyzomyia Listeri *sergentii Cellia* Myzomyia Demeilloni *seydeli Cellia* Myzomyia Marshallii

*rufipes Cellia* Neocellia

*salbaii Cellia* Neocellia

*saungi Cellia* Neomyzomyia

*schwetzi Cellia* Myzomyia

*shannoni Anopheles* Laticorn Arribalzagia

*rodhaini Cellia rollai Kerteszia*

34 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors


**Species Subgenus Section Series Group Subgroup**

*trinkae Nyssorhynchus* Albimanus Oswaldoi Oswaldoi Oswaldoi

*umbrosus Anopheles* Laticorn Myzorhynchus Umbrosus Umbrosus

*vaneedeni Cellia* Myzomyia Funestus Funestus

*vanus Anopheles* Laticorn Myzorhynchus Barbirostris Vanus

*varuna Cellia* Myzomyia Funestus Aconitus

*vietnamensis Anopheles* Laticorn Myzorhynchus Hyrcanus Lesteri

*whartoni Anopheles* Laticorn Myzorhynchus Umbrosus Letifer

*yaeyamaensis Cellia* Myzomyia Funestus Minimus

*tibiamaculatus Anopheles* Angusticorn Anopheles Pseudopunctipennis

*tigertti Anopheles* Angusticorn Anopheles Aitkenii *torresiensis Cellia* Neomyzomyia Punctulatus *triannulatus Nyssorhynchus* Albimanus Oswaldoi Triannulatus

36 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

*turkhudi Cellia* Paramyzomyia Cinereus

*vagus Cellia* Pyretophorus Subpictus

*vanhoofi Cellia* Neomyzomyia Smithii

*vernus Cellia* Neomyzomyia Ardensis

*vinckei Cellia* Neomyzomyia Ardensis *walkeri Anopheles* Angusticorn Anopheles Maculipennis

*wellcomei Cellia* Myzomyia Wellcomei *wellingtonianus Anopheles* Angusticorn Anopheles Lindesayi

*willmori Cellia* Neocellia Maculatus *wilsoni Cellia* Neomyzomyia Smithii *xelajuensis Anopheles* Angusticorn Anopheles Plumbeus *xui Anopheles* Laticorn Myzorhynchus Hyrcanus

*ziemanni Anopheles* Laticorn Myzorhynchus Coustani

*veruslanei Anopheles* Laticorn Arribalzagia *vestitipennis Anopheles* Laticorn Arribalzagia

*walravensi Cellia* Myzomyia *watsonii Cellia* Neomyzomyia

*vargasi Lophopodomyia*

Sibling species complexes of *Anopheles* – formally named and unnamed species. The Macula‐ tus, Maculipennis and Punctulatus Complexes are now considered to be super-complexes referred to as "Groups" with subordinate complexes. Likewise, the Culicifacies Complex is considered to be a Subgroup. Appendix 3. Sibling species complexes of Anopheles– formally named and unnamed species. The Maculatus, Maculipennis and Punctulatus Complexes are now considered to be super-complexes referred to as "Groups" with subordinate complexes. Likewise,

the Culicifacies Complex is considered to be a Subgroup. Subgenus Anopheles Barbirostris Complex [76] barbirostris Claviger Complex [67] claviger petragnani Crucians Complex [72] bradleyi crucians A crucians B crucians C crucians D crucians E georgianus Gigas Complex [70] baileyi gigas gigas s.l. Lindesayi Complex [70] lindesayi mengalangensis nilgiricus wellingtonianus Maculipennis Group [20] atropos aztecus lewisi walkeri Maculipennis Subgroup [71] artemievi atroparvus daciae labranchiae maculipennis martinius melanoon messeae persiensis sacharovi Quadrimaculatus Subgroup [21] beklemishevi diluvialis inundatus maverlius quadrimaculatus smaragdinus Freeborni Subgroup [71] earlei freeborni hermsi occidentalis

 Subgenus Cellia Annularis Complex [86] annularis A annularis B pallidus philippinensis schueffneri Annulipes Complex [89] annulipes A annulipes B annulipes C annulipes D annulipes F annulipes G annulipes H annulipes I annulipes J annulipes K annulipes L annulipes M annulipes N annulipes O annulipes P annulipes Q Culicifacies Subgroup [108] culicifacies A culicifacies B culicifacies C culicifacies D culicifacies E Dirus Complex [93] aff. takasagoensis baimaii cracens dirus elegans nemophilous scanloni takasagoensis Fluviatilis Complex [83] fluviatilis S fluviatilis T fluviatilis U Gambiae Complex [96] arabiensis bwambae comorensis gambiae melas merus quadriannulatus quadriannulatus B Leucosphyrus Complex [91] baisasi balabacensis introlatus

 latens leucosphyrus Longirostris Complex [90] Genotype A Genotype B Genotype C1 Genotype C2 Genotype D Genotype E Genotype F Genotype G Genotype H Lungae Complex [47] lungae nataliae solomonis Maculatus Group [88] dispar greeni pseudowillmori willmori Maculatus Subgroup [73] dravidicus maculatus Sawadwongporni Subgroup [73] notanandai rampae sawadwongporni Marshallii Complex [85] hughi kosiensis letabensis marshallii Minimus Complex [84] harrisoni minimus yaeyamaensis Nivipes Complex [70,87] nivipes cytotype 1 nivipes cytotype 2 Nili Complex [22] carnevalei nili ovengensis somalicus Punctulatus Group [94] clowi koliensis punctulatus rennellensis sp. nr punctulatus Farauti Complex [94,109] farauti farauti 4

farauti 5 farauti 6 farauti 8 hinesorum irenicus torresiensis Subpictus Complex [95] subpictus A subpictus B subpictus C subpictus D Sundaicus Complex [97] epiroticus sundaicus sundaicus B sundaicus C sundaicus D sundaicus E Superpictus Complex [110] superpictus A superpictus B Subgenus Kerteszia Cruzii Complex [101] cruzii A cruzii B cruzii C Subgenus Nyssorhynchus Albitarsis Complex [106] albitarsis albitarsis F albitarsis G albitarsis H albitarsis I deaneorum janconnae lineage nr janconnae marajoara oryzalimnetes Benarrochi Complex [105] benarrochi benarrochi B Nuneztovari Complex [104,111] goeldii nuneztovari B/C nuneztovari A Triannulatus Complex [112,113] halophylus triannulatus triannulatus C

Unnamed and provisionally designated members of species complexes and their position in the classification of genus *Anopheles* (Sections of subgenera *Anopheles* and *Nyssorhynchus* are omitted). Excluding nominotypical members, the list includes 72 species that require formal Latin names.



**Appendix 4**

38 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Latin names.

*cruzii* sp. A,B,C [118] *Kerteszia*

*longirostris* Genotypes A,B,C1,C2,D,E,F,G,H

Unnamed and provisionally designated members of species complexes and their position in the classification of genus *Anopheles* (Sections of subgenera *Anopheles* and *Nyssorhynchus* are omitted). Excluding nominotypical members, the list includes 72 species that require formal

**Species Authors Subgenus Series Group Subgroup Complex** *albitarsis* sp. F,G,H,I [114,115] *Nyssorhynchus* Albitarsis Albitarsis Albitarsis *annularis* sp. A,B [86] *Cellia* Neocellia Annularis Annularis *annulipes* sp. A‒Q [89] *Cellia* Neomyzomyia Annulipes

*barbirostris* clades I‒IV [117] *Anopheles* Barbirostris Barbirostris Barbirostris Barbirostris *benarrochi* sp. B [105] *Nyssorhynchus* Oswaldoi Oswaldoi Strodei Benarrochi *crucians* sp. A‒E [72] *Anopheles* Anopheles Punctipennis Crucians

*culicifacies* sp. A‒E [108] *Cellia* Myzomyia Funestus Culicifacies Culicifacies *farauti* sp. 4,5,6 [109,119] *Cellia* Neomyzomyia Punctulatus Farauti *fluviatilis* sp. S,T,U [83] *Cellia* Myzomyia Funestus Minimus Fluviatilis

*gigas s.l.* (Thailand) [70] *Anopheles* Anopheles Lindesayi Gigas

*janconnae*, lineage nr [122] *Nyssorhynchus* Albitarsis Albitarsis Albitarsis

*marajoara* lineages 1,2 [124] *Nyssorhynchus* Albitarsis Albitarsis Albitarsis *nivipes* (2 cytotypes) [87] *Cellia* Neocellia Annularis Nivipes *nuneztovari* sp. A [125] *Nyssorhynchus* Oswaldoi Oswaldoi Oswaldoi Nuneztovari *nuneztovari* B/C [104] *Nyssorhynchus* Oswaldoi Oswaldoi Oswaldoi Nuneztovari

[90] Cellia Neomyzomyia Longirostris

*Anopheles* CP Form [116] *Nyssorhynchus* Oswaldoi Oswaldoi Strodei

*funestus*-like sp. [120] *Cellia* Myzomyia Funestus Funestus

*longipalpis* Type A [123] *Cellia* Myzomyia Funestus Minimus *longipalpis* Type C [123] *Cellia* Myzomyia Funestus Funestus

*hyrcanus* spIR [121] *Anopheles* Hyrcanus

*punctulatus*, sp. nr [126] *Cellia* Neomyzomyia Punctulatus

Phylogenetic studies of *Anopheles* mosquitoes. Groups included in the table are those recog‐ nized herein. None of the studies included all taxa that comprise the group investigated, but those marked with an asterisk (\*) included the majority of species. Nucleotide sequences include *COI*, *COII*, *cyt b*, *ND4*, *ND5* and *ND6* from mitochondrial DNA (mtDNA); *D2*, *D3*, *18S*, *ITS1* and *ITS2* from ribosomal DNA (rDNA); *EF-1α*, *G6pd* and *white* from nuclear DNA.



**Group Data set Authors**

Myzomyia Series Chromosomes [143,144]

*COII, D3*

*COII*, *D3 D3, ITS2*

*D3, ITS2*

Maculatus Group *ITS2*, *COII*, *D3* [149–151]

*COI, ND6*

*18S*

Farauti Complex *ITS1* [109]

Gambiae Complex Chromosomes [155]

*white cyt b, ITS2, COI*

Subgenus *Kerteszia* Morphology [44] Subgenus *Nyssorhynchus ITS2* [159]

Albimanus Section Morphology [24] Argyritarsis Section Morphology [25] Myzorhynchella Section *ITS2*, *COI*, *white* [160]

Chromosomes

*ITS2*, *D2*, *COI*, *ND4*

*COII* [37]

rDNA, mtDNA [156]

Chromosomes *ITS2*, *COII*, *D3*, *ND5*

*ITS2*, *COI*, *COII*, EF-1α

Subgenus *Cellia* Chromosomes [143,144]

Funestus Group *ITS2, COII, D3*

40 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Punctulatus Group *ITS2*

Pyretophorus Series Morphology

Sundaicus Complex mtDNA

Minimus Subgroup Minimus Complex

Neocellia Series Annularis Group

Neomyzomyia Series Annulipes Complex Leucosphyrus Group *ITS2*, *COI* [142]

*COII* [37]

*COII*, *D3* [82]

[55]\* [81]\*

[145] [146]\*

[87] [147] [148]

[89] [152]

[153] [154]

[56]\* [144]

[157] [158] [114] [128] Summary of the formal and informal group taxa (species complexes omitted) of genus *Anopheles*. The zoogeographic distribution and the number of formally named and informally designated species (in parentheses) are given for each taxon. Minor extensions of one or more species of a group into an adjacent zoogeographic region are disregarded. C = cosmopolitan; NW = New World; OW = Old World; Af = Afrotropical; Au = Australasian; Ne = Nearctic; Nt = Neotropical; Or = Oriental; Pa = Palaearctic. Appendix 6. Summary of the formal and informal group taxa (species complexes omitted) of genus Anopheles. The zoogeographic

distribution and the number of formally named and informally designated species (in parentheses) are given for each taxon. Minor extensions of one or more species of a group into an adjacent zoogeographic region are disregarded. C = cosmopolitan; NW = New World; OW = Old World; Af = Afrotropical; Au = Australasian; Ne = Nearctic; Nt = Neotropical; Or = Oriental; Pa = Palaearctic.

Subgenus Anopheles‒ C (191) Angusticorn Section ‒ OW, NW (95) Anopheles Series ‒ OW and NW (88) Alongensis Group ‒ Or (2) Aitkenii Group ‒ Or (13) Atratipes Group ‒ Au (2) Culiciformis Group ‒ Or (3) Lindesayi Group ‒ Or (7) Maculipennis Group ‒ Ne, Pa (20) Maculipennis Subgroup ‒ Pa (10) Quadrimaculatus Subgroup ‒ Ne (5), Pa (1) Freeborni Subgroup ‒ Ne (4) Plumbeus Group ‒ Ne (2), Nt (4), Pac (3) Pseudopunctipennis Group ‒ Ne (7) Punctipennis Group ‒ Ne (9) Stigmaticus Group ‒ Au (6) Cycloleppteron Series ‒ Ne (2) Lophoscelomyia Series ‒ Or (4) Unassigned ‒ (1) Asiaticus Group ‒ (4) Unassigned ‒ (2) Asiaticus Subgroup ‒ (1) Interruptus Subgroup ‒ (1) Laticorn Section ‒ Af, Au, Nt, Or, Pa (96) Arribalzagia Series ‒ Ne (24) Christya Series ‒ Af (2) Myzorhynchus Series ‒ Af, Au, Or, Pa (70) Albotaeniatus Group ‒ Or (4), Pa (1) Bancroftii Group ‒ Au/Or (2) Barbirostris Group ‒ Or (16) Unassigned ‒ (2) Barbirostris Subgroup ‒ (9) Vanus Subgroup ‒ (5) Coustani Group ‒ Af (9) Hyrcanus Group ‒ Pa (26) Unassigned ‒ (17) Lesteri Subgroup ‒ (5) Nigerrimus Subgroup ‒ (4)

 Umbrosus Group ‒ Or (12) Unassigned ‒ (5) Baezai Subgroup ‒ (1) Letifer Subgroup ‒ (4) Separatus Subgroup ‒ (1) Umbrosus Subgroup ‒ (1)

Subgenus Baimaia‒ Or (1)

Subgenus Cellia‒ OW (233) Cellia Series ‒ Af (8) Unassigned ‒ (6) Squamosus Group ‒ (2) Myzomyia Series ‒ Af, Or (71) Unassigned ‒ Af (16) Demeilloni Group ‒ Af (7) Funestus Group ‒ Af, Or (29) Unassigned ‒ (1) Aconitus Subgroup ‒ Or (5) Culicifacies Subgroup ‒ Or (5) Funestus Subgroup ‒ Af (7) Minimus Subgroup ‒ Af (1), Or (6) Rivulorum Subgroup ‒ Af (4) Marshallii Group ‒ Af (15) Wellcomei Group ‒ Af (4) Neocellia Series ‒ Af, Or, Pal (24) Unassigned ‒ Af, Or, Pa (14) Annularis Group ‒ Or (7) Jamesii Group ‒ Or (3) Maculatus Group ‒ Or (9) Unassigned ‒ (4) Maculatus Subgroup ‒ (2) Sawadwongporni Subgroup ‒ (3) Neomyzomyia Series ‒ Af, Au, Or (121) Unassigned ‒ Af, Au, Or (42) Ardensis Group ‒ Af (18) Kochi Group ‒ Or (1) Leucosphyrus Group ‒ O (21) Hackeri Subgroup ‒ (5) Leucosphyrus Subgroup ‒ (13) Riparis Subgroup ‒ (3)

Mascarensis Group ‒ Af (1) Pauliani Group ‒ Af (5) Punctulatus Group ‒ Au (13) Ranci Group ‒ Af (5) Unassigned ‒ (1) Ranci Subgroup ‒ (1) Roubaudi Subgroup ‒ (3) Rhodesiensis Group ‒ Af (5) Smithii Group ‒ Af (9) Tessellatus Group ‒ Or (1) Paramyzomyia Series ‒ Af, Pa (6) Cinereus Group ‒ Af (2), Pa (1) Listeri Group ‒ Af (2), Pa (1 Pyretophorus Series ‒ Af (10), Or (12)

Subgenus Kerteszia‒ Nt (14)

Subgenus Lophopodomyia‒ Nt (6)

Subgenus Nyssorhynchus‒ Nt (38) Albimanus Section ‒ (24) Albimanus Series ‒ (1) Oswaldoi Series ‒ (23) Oswaldoi Group ‒ (21) Oswaldoi Subgroup ‒ (14) Strodei Subgroup ‒ (7) Triannulatus Group ‒ (3) Argyritarsis Section ‒ (14) Albitarsis Series ‒ (8) Albitarsis Group ‒ (7) Braziliensis Group ‒ (1) Argyritarsis Series ‒ (6) Argyritarsis Group ‒ (2) Darlingi Group ‒ (1) Lanei Group ‒ (1) Pictipennis Group ‒ (2) Myzorhynchella Section ‒ (6)

Subgenus Stethomyia‒ Ne (5)

## **Author details**

Ralph E. Harbach\*

Address all correspondence to: r.harbach@nhm.ac.uk

Department of Life Sciences, Natural History Museum, London, UK

Three new species of *Anopheles* were formally described and named while the book was in press: *An*. (*Anopheles*) *vanderwulpi* (= *An. barbirostris* clade II) [161]; *An*. (*Cellia*) *amharicus* (= *An. quadriannulatus* sp. B) and *An*. (*Cellia*) *coluzzii* (= molecular M form of *An. gambiae*) [162]. *Anopheles* (*Anopheles*) *kunmingensis* (Laticorn Section, Myzorhynchus Series, Hyrcanus Group) was inadvertently omitted from Appendix 2 during preparation of the chapter. Thus, the genus now includes 469 formally named species and 70 species that require formal Latin names.

## **References**


[10] Edwards FW. A revision of the mosquitos [*sic*] of the Palaearctic Region. Bulletin of Entomological Research 1921;12 263–351.

**Author details**

Ralph E. Harbach\*

Latin names.

**References**

Address all correspondence to: r.harbach@nhm.ac.uk

42 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Insekten. Volume 1. Aachen; 1818.

ish Museum (Natural History); 1901.

ish Museum (Natural History); 1901.

ish Museum (Natural History); 1903.

ish Museum (Natural History); 1907.

ish Museum (Natural History); 1910.

1915;3 371–394.

Second edition. Thomas Say Foundation 1977;6 1–611.

Tropical Diseases and Preventive Medicine 1913;1 33–43.

Department of Life Sciences, Natural History Museum, London, UK

Three new species of *Anopheles* were formally described and named while the book was in press: *An*. (*Anopheles*) *vanderwulpi* (= *An. barbirostris* clade II) [161]; *An*. (*Cellia*) *amharicus* (= *An. quadriannulatus* sp. B) and *An*. (*Cellia*) *coluzzii* (= molecular M form of *An. gambiae*) [162]. *Anopheles* (*Anopheles*) *kunmingensis* (Laticorn Section, Myzorhynchus Series, Hyrcanus Group) was inadvertently omitted from Appendix 2 during preparation of the chapter. Thus, the genus now includes 469 formally named species and 70 species that require formal

[1] Meigen JW. Systematische Beschreibung der bekannten europäischen zweiflügeligen

[2] Theobald FV. A Monograph of the Culicidae or Mosquitoes. Volume 1. London: Brit‐

[3] Theobald FV. A Monograph of the Culicidae or Mosquitoes. Volume 2. London: Brit‐

[4] Theobald FV. A Monograph of the Culicidae or Mosquitoes. Volume 3. London: Brit‐

[5] Theobald FV. A Monograph of the Culicidae or Mosquitoes. Volume 4. London: Brit‐

[6] Theobald FV. A Monograph of the Culicidae or Mosquitoes. Volume 5. London: Brit‐

[7] Knight KL, Stone A. A catalog of the mosquitoes of the world (Diptera: Culicidae).

[8] Knab F. The species *of Anopheles* that transmit human malaria. American Journal of

[9] Christophers SR. The male genitalia of *Anopheles*. Indian Journal of Medical Research


[38] Krzywinski J, Wilkerson RC, Besansky N. Evolution of mitochondrial and ribosomal gene sequences in Anophelinae (Diptera: Culicidae): implications for phylogeny re‐ construction. Molecular Phylogenetics and Evolution 2001;18 479–487.

[25] Linthicum KJ. A revision of the Argyritarsis Section of the subgenus *Nyssorhynchus* of

[26] Harbach RE. Review of the internal classification of the genus *Anopheles* (Diptera: Cu‐ licidae): the foundation for comparative systematics and phylogenetic research. Bul‐

[27] Harbach RE. The classification of genus *Anopheles* (Diptera: Culicidae): a working hy‐ pothesis of phylogenetic relationships. Bulletin of Entomological Research 2004;94

[28] Peyton EL, Wilkerson RC, Harbach RE. Comparative analysis of the subgenera *Ker‐ teszia* and *Nyssorhynchus* of *Anopheles* (Diptera: Culicidae). Mosquito Systematics

[29] Borkent A, Grimaldi DA. The earliest fossil mosquito (Diptera: Culicidae), in mid-Cretaceous amber. Annals of the Entomological Society of America 2004;97 882–888.

[30] Bertone MA, Courtney GW, Wiegmann BM. Phylogenetics and temporal diversifica‐ tion of the earliest true flies (Insecta: Diptera) based on multiple nuclear genes. Sys‐

[31] Poinar Jr JO, Zavortink TJ, Pike T, Johnston PA. *Paleoculicis minutus* (Diptera: Culici‐ dae) n. gen., n. sp., from Cretaceous Canadian amber, with a summary of described

[32] Zavortink TJ, Poinar Jr GO. *Anopheles* (*Nyssorhynchus*) *dominicanus* sp. n. (Diptera: Culicidae) from Dominican amber. Annals of the Entomological Society of America

[33] Statz G. Neue Dipteren (Nematocera) aus dem Oberoligocän von Rott. II. Teil. V. Familie Culicidae (Stechmücken). Palaeontographica 1944;95(A) 108–120, 6 pls.

[34] Reidenbach KR, Cook S, Bertone MA, Harbach RE, Wiegmann BM, Besansky NJ. Phylogenetic analysis and temporal diversification of mosquitoes (Diptera: Culici‐ dae) based on nuclear genes and morphology. BMC Evolutionary Biology 2009;9 298.

[35] Besansky NJ, Fahey GT. Utility of the *white* gene in estimating phylogenetic relation‐ ships among mosquitoes (Diptera: Culicidae). Molecular Biology and Evolution

[36] Harbach RE, Kitching IJ. Reconsideration of anopheline phylogeny (Diptera: Culici‐ dae: Anophelinae) based on morphological data. Systematics and Biodiversity 2005;3

[37] Foley DH, Bryan JH, Yeates D, Saul A. Evolution and systematics of *Anopheles*: in‐ sights from a molecular phylogeny of Australasian mosquitoes. Molecular Phyloge‐

fossil mosquitoes. Acta Geologica Hispanica 2000;35 119–128.

*Anopheles* (Diptera: Culicidae). Mosquito Systematics 1988;20 98–271.

letin of Entomological Research 1994;84 331–342.

tematic Entomology 2008;33 668‒687.

537–553.

1992;24 51–69.

44 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

2000;93 1230–1235.

1997;14 442–454.

netics and Evolution 1998;9 262–275.

345–374.


[64] Thelwell NJ, Huisman RA, Harbach RE, Butlin RK. Evidence for mitochondrial intro‐ gression between *Anopheles bwambae* and *Anopheles gambiae*. Insect Molecular Biology 2000;9 203–210.

[52] Mattingly PF. The Biology of Mosquito-Borne Disease. The Science of Biology Series

[53] Sinka ME, Bangs MJ, Manguin S, Coetzee M, Mbogo CM, Hemingway J, Patil AP, Temperley WH, Gething PW, Kabaria CW, Okara RM, Van Boeckel T, Godfray HCJ, Harbach RE, Hay SI. The dominant *Anopheles* vectors of human malaria in Africa, Eu‐ rope and the Middle East: occurrence data, distribution maps and bionomic précis.

[54] Sinka ME, Bangs MJ, Manguin S, Chareonviriyaphap T, Patil AP, Temperley WH, Gething PW, Elyazar IRF, Kabaria CW, Harbach RE, Hay SI. The dominant *Anopheles* vectors of human malaria in the Asia-Pacific region: occurrence data, distribution

[55] Garros C, Harbach RE, Manguin S. Systematics and biogeographical implications of the phylogenetic relationships between members of the Funestus and Minimus Groups of *Anopheles* (Diptera: Culicidae). Journal of Medical Entomology 2005;42 7–

[56] Anthony TG, Harbach RE, Kitching IJ. Phylogeny of the Pyretophorus Series of *Anopheles* subgenus *Cellia* (Diptera: Culicidae). Systematic Entomology 1999;24 193–

[57] Evans AM. Mosquitoes of the Ethiopian Region. II.–Anophelini adults and early

[58] Gillies MT. Notes on the eggs of some East African *Anopheles.* Annals of Tropical

[61] Sinka ME, Rubio-Palis Y, Manguin S, Patil AP, Temperley WH, Gething PW, Van Boeckel T, Kabaria CW, Harbach R.E, Hay SI. The dominant *Anopheles* vectors of hu‐ man malaria in the Americas: occurrence data, distribution maps and bionomic pré‐

[62] Sinka ME, Bangs MJ, Manguin S, Rubio-Palis Y, Chareonviriyaphap T, Coetzee M, Mbogo CM, Hemingway J, Patilm AP, Temperley WH, Gething PW, Kabaria CW, Okara RM, Burkot TR, Harbach RE, Hay SI. A global map of dominant malaria vec‐

[63] Besansky NJ, Lehmann T, Fahey GT, Fontenille D, Braack LEO, Hawley WA, Collins FH. Patterns of mitochondrial variation within and between African malaria vectors, *Anopheles gambiae* and *An. arabiensis*, suggest extensive gene flow. Genetics 1997;147

[59] Mattingly PF. Mosquito eggs III. Mosquito Systematics Newsletter 1969;1 41–50.

[60] Stringer C. The Origin of our Species. London: Penguin Books Ltd; 2011.

1. London: George Allen & Unwin Ltd; 1969.

maps and bionomic précis. Parasites & Vectors 2011;4 89.

stages. London: British Museum (Natural History); 1938.

Medicine and Parasitology 1955;49 158–160.

cis. Parasites & Vectors 2010;3 72.

tors. Parasites & Vectors 2012;5 69.

1817‒1828.

Parasites & Vectors 2010;3 117.

46 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

18.

205.


[89] Foley DH, Wilkerson RC, Cooper RD, Volovsek ME, Bryan JH. A molecular phyloge‐ ny of *Anopheles annulipes* (Diptera: Culicidae) sensu lato: The most species-rich ano‐ pheline complex. Molecular Phylogenetics and Evolution 2007;43 283–297.

[76] Satoto TBT. Cryptic species within *Anopheles barbirostris* van der Wulp, 1884, inferred from nuclear and mitochondrial gene sequence variation. PhD Thesis. University of

[77] Reid JA. The *Anopheles hyrcanus* group in south–east Asia (Diptera: Culicidae). Bulle‐

[78] Harrison BA. A new interpretation of affinities within the *Anopheles hyrcanus* com‐

[79] Reid JA. The *Anopheles umbrosus* group (Diptera: Culicidae). Part 1: systematics, with descriptions of two new species. Transactions of the Royal Entomological Society of

[80] Theobald FV. The classification of the Anophelina. Journal of Tropical Medicine

[81] Garros C, Harbach RE, Manguin S. Morphological assessment and molecular phylo‐ genetics of the Funestus and Minimus Groups of *Anopheles* (*Cellia*). Journal of Medi‐

[82] Chen B, Butlin RK, Harbach RE. Molecular phylogenetics of the Oriental members of the Myzomyia Series of *Anopheles* subgenus *Cellia* (Diptera: Culicidae) inferred from nuclear and mitochondrial DNA sequences. Systematic Entomology 2003;28 57–69.

[83] Sarala KS, Nutan N, Vasantha K, Dua VK, Malhotra MS, Yadav RS, Sharma VP. Cy‐ togenetic evidence for three sibling species in *Anopheles fluviatilis* (Diptera: Culici‐

[84] Green CA, Gass RF, Munstermann LE, Baimai V. Population-genetic evidence for two species in *Anopheles minimus* in Thailand. Medical and Veterinary Entomology

[85] Gillies MT, Coetzee M. A supplement to the Anophelinae of Africa south of the Sa‐ hara (Afrotropical Region). Publications of the South African Institute for Medical

[86] Atrie B, Subbarao SK, Pillai MKK, Rao SRV, Sharma VP. Population cytogenetic evi‐ dence for sibling species in *Anopheles annularis* (Diptera: Culicidae). Annals of the En‐

[87] Green CA, Harrison BA, Klein TA, Baimai V. Cladistic analysis of polytene chromo‐ some rearrangements in anopheline mosquitoes, subgenus *Cellia*, series *Neocellia*.

[88] Rattanarithikul R, Green CA. Formal recognition of the species of the *Anopheles macu‐ latus* group (Diptera: Culicidae) occurring in Thailand, including the descriptions of two new species and a preliminary key to females. Mosquito Systematics 1987;18

dae). Annals of the Entomological Society of America 1994;87 116–121.

Liverpool; 2001.

48 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

London 1950;101 281–318.

cal Entomology 2005;42 522–536.

1902;5 181–183.

1990;4 25–34.

246–278.

Research 1987;55 1–143.

tomological Society of America 1999;92 243–249.

Canadian Journal of Genetics and Cytology 1985;27 123–133.

tin of Entomological Research 1953;44 5–76.

plex of Southeast Asia. Mosquito Systematics 1972;4 73–83.


[113] Silva-do-Nascimento TR, Lourenço-de-Oliveira R. Diverse population dynamics of three *Anopheles* species belonging to the Triannulatus Complex (Diptera: Culicidae). Memorias do Instituto Oswaldo Cruz 2007;102 975–982.

[101] Ramirez CC, Dessen EM. Chromosomal evidence for sibling species of the malaria

[102] Blanchard R. Nouvelle note sur les moustiques. Comptes Rendus Hebdomadaires

[103] Levi Castillo R. Atlas de los Anofelinos Sudamericanos. Quayaquil, Ecuador: Socie‐

[104] Sierra DM, Velez ID, Linton Y-M. Malaria vector *Anopheles* (*Nyssorhynchus*) *nunezto‐ vari* comprises one genetic species in Colombia based on homogeneity of nuclear

[105] Ruiz F, Quiñones ML, Erazo HF, Calle DA, Alzate JF, Linton Y-M. Molecular differ‐ entiation of *Anopheles* (*Nyssorhynchus*) *benarrochi* and *An.* (*N.*) *oswaldoi* from Southern

[106] Wilkerson RC, Parsons TJ, Klein TA, Gaffigan TV, Bergo E, Consolim J. Diagnosis by random amplified polymorphic DNA polymerase chain reaction for four cryptic spe‐ cies related to *Anopheles* (*Nyssorhynchus*) *albitarsis* (Diptera: Culicidae) from Para‐

guay, Argentina, and Brazil. Journal of Medical Entomology 1995;32 697–704. [107] Peyton EL, Wilkerson RC, Harbach RE. Comparative analysis of the subgenera *Ker‐ teszia* and *Nyssorhynchus* of *Anopheles* (Diptera: Culicidae). Mosquito Systematics

[108] Kar I, Subbarao SK, Eapen A, Ravendaran J, Satyanarayana TS, Raghavendra K, Nan‐ da N, Sharma VP. Evidence for a new malaria vector species, species E, within the *Anopheles culicifacies* complex (Diptera: Culicidae). Journal of Medical Entomology

[109] Bower JE, Dowton M, Cooper RD, Beebe NW. Intraspecific concerted evolution of the rDNA ITS1 in *Anopheles farauti* sensu stricto (Diptera: Culicidae) reveals recent patterns of population structure. Journal of Molecular Evolution 2008;67 397‒411.

[110] Oshaghi MA, Yaghobi-Ershadi MR, Shemshad Kh, Pedram M, Amani H. The *Anoph‐ eles superpictus* complex: introduction of a new malaria vector complex in Iran. Bulle‐

[111] Conn J, Puertas YR, Seawright JA. A new cytotype *of Anopheles nuneztovari* from western Venezuela and Colombia. Journal of the American Mosquito Control Associ‐

[112] Rosa-Freitas MG, Lourenço-de-Oliveira R, de Carvalho-Pinto CJ, Flores-Mendoza F, Silva-do-Nascimento TF. Anopheline species complexes in Brazil. Current knowl‐ edge of those related to malaria transmission. Memorias do Instituto Oswaldo Cruz

tin de la Société de Pathologie exotique 2008;101 429‒434.

des Séances et Mémoires de la Société de Biologie 1902;54 793–795.

ITS2 rDNA. Journal of Medical Entomology 2004;41 302–307.

Colombia. Memorias do Instituto Oswaldo Cruz 2005;100 155–160.

vector *Anopheles cruzii*. Genome 2000;43 143–151.

dad Filantrópica de Guayas; 1949.

50 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

1992;24 51–69.

1999;36 595–600.

ation 1993;9 294–301.

1998;93 651–655.


[136] Porter CH, Collins FH. Phylogeny of Nearctic members of the *Anopheles maculipennis* species group derived from the D2 variable region of 28S ribosomal RNA. Molecular Phylogenetics and Evolution 1996;6 178–188.

[124] McKeon SN, Lehr MA, Wilkerson RC, Ruiz JF, Sallum MA, Povoa MM., Conn JE, Li‐ ma JBP. Lineage divergence detected in the malaria vector *Anopheles marajoara* (Dip‐

[125] Conn J. A genetic study of the malaria vector *Anopheles nuneztovari* from western Venezuela. Journal of the American Mosquito Control Association 1990;6 400–405.

[126] Foley DH, Cooper RD, Bryan JH. A new species within the *Anopheles punctulatus* complex in Western Province, Papua New Guinea. Journal of the American Mosqui‐

[127] Hunt RH, Coetzee M, Fettene M. The *Anopheles gambiae* complex: a new species from Ethiopia. Transactions of the Royal Society of Tropical Medicine and Hygiene

[128] Dusfour I, Michaux JR, Harbach RE, Manguin S. Speciation and phylogeography of the Southeast Asian *Anopheles sundaicus* complex. Infection, Genetics and Evolution

[129] Oshaghi MA, Shemshad Kh, Yaghobi-Ershadi MR, Pedram M, Vatandoost H, Abaie MR, Akbarzadeh K, Mohtarami F. Genetic structure of the malaria vector *Anopheles superpictus* in Iran using mitochondrial cytochrome oxidase (COI and COII) and mor‐

[130] Takano KT, Nguyen NTH, Nguyen BTH, Sunahara T, Yasunami, M, Nguyen MD, Takagi M. Partial mitochondrial DNA sequences suggest the existence of a cryptic species within the Leucosphyrus group [*sic*] of the genus *Anopheles* (Diptera: Culici‐ dae), forest malaria vectors, in northern Vietnam. Parasites & Vectors 2010;3 41.

[131] Bargues MD, Latorre JM, Morchon R, Simon F, Escosa R, Aranda C, Sainz S, Fuentes MV, Mas-Coma S. rDNA sequences of *Anopheles* species from the Iberian Peninsula and an evaluation of the 18S rRNA gene as phylogenetic marker in Anophelinae.

[132] White GB. Systematic reappraisal of the *Anopheles maculipennis* complex. Mosquito

[133] Gordeev M, Goriacheva I, Shaikevitch E, Ejov M. Variability of the second internal transcribed spacer of the ribosomal DNA among five Palaearctic species of anophe‐

[134] Djadid ND, Gholizadeh S, Tafsiri E, Romi R, Gordeev M, Zakeri S. Molecular identi‐ fication of Palearctic [*sic*] members of *Anopheles maculipennis* in northern Iran. Malar‐

[135] Marinucci M, Romi R, Mancini M, Di Luca M, Severini C. Phylogenetic relationships of seven palearctic [*sic*] members of the *maculipennis* complex inferred from ITS2 se‐

Journal of Medical Entomology 2006;43 508–517.

line mosquitoes. European Mosquito Bulletin 2004;17 14–19.

quence analysis. Insect Molecular Biology 1999;8 469480.

Systematics 1978;10 13–44.

ia Journal 2007;6-6 (10 pp.).

phologic markers: A new species complex? Acta Tropica 2007;101 241‒248.

tera: Culicidae) in Amazonian Brazil. Malaria Journal 2010;9 271.

to Control Association 1995;11 122–127.

52 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

1998;92 231–235.

2007;7 484‒493.


netics and biogeography of the Neocellia Series of *Anopheles* mosquitoes in the Orien‐ tal Region. Molecular Phylogenetics and Evolution 2009;52 588–601.


[159] Marrelli MT, Floeter-Winter LM, Malafronte RS, Tadei WP, Lourenço-de-Oliveira R, Flores-Mendoza C, Marinotti O. Amazonian malaria vector anopheline relationships interpreted from ITS2 rDNA sequences. Medical and Veterinary Entomology 2005;19 208–218.

netics and biogeography of the Neocellia Series of *Anopheles* mosquitoes in the Orien‐

[148] Alam MT, Das MK, Dev V, Ansari MA, Sharma YD. PCR-RFLP method for the iden‐ tification of four members of the *Anopheles annularis* group of mosquitoes (Diptera: Culicidae). Transactions of the Royal Society of Tropical Medicine and Hygiene

[149] Ma Y, Qu F, Dong X, Zhou H. Molecular identification of *Anopheles maculatus* com‐ plex from China. Chinese Journal of Parasitology and Parasitic Diseases 2002;20 321–

[150] Ma Y, Shizhu L, Jiannong X. Molecular identification and phylogeny of the Macula‐ tus Group of *Anopheles* mosquitoes (Diptera: Culicidae) based on nuclear and mito‐

[151] Walton C, Somboon P, Harbach RE, Zhang S, Weerasinghe I, O'Loughlin SM, Phom‐ pida S, Sochantha T, Tun-Lin W, Chen B, Butlin RK. Molecular identification of mos‐ quito species in the *Anopheles annularis* group in southern Asia. Medical and

[152] Sallum MAM, Foster PG, Li C, Sithiprasasna R, Wilkerson RC. Phylogeny of the Leu‐ cosphyrus Group of *Anopheles* (*Cellia*) Diptera: Culicidae) based on mitochondrial gene sequences. Annals of the Entomological Society of America 2007;100 27–35.

[153] Beebe NW, Ellis JT, Cooper RD, Saul A. DNA sequence analysis of the ribosomal DNA ITS2 region for the *Anopheles punctulatus* group of mosquitoes. Insect Molecular

[154] Beebe NW, Cooper RE. Distribution and evolution of the *Anopheles punctulatus* group (Diptera: Culicidae) in Australia and Papua New Guinea. International Journal of

[155] Coluzzi M, Sabatini A, Petrarca V, Di Deco MA. Chromosomal differentiation and adaptation to human environments in the *Anopheles gambiae* complex. Transactions of

[156] Besansky NJ, Powell JR, Caccone A, Hamm DM, Scott JA, Collins FH. Molecular phylogeny of the *Anopheles gambiae* complex suggests genetic introgression between principal malaria vectors. Proceedings of the National Academy of Science, USA

[157] Caccone A, Garcia BA, Powell JR. Evolution of the mitochondrial DNA control re‐ gion in the *Anopheles gambiae* complex. Insect Molecular Biology 1996;6 51–59.

[158] Wilkerson RC, Foster PG, Li C, Sallum MAM. Molecular phylogeny of Neotropical *Anopheles* (*Nyssorhynchus*) *albitarsis* species complex (Diptera: Culicidae). Annals of

the Entomological Society of America 2005;98 918–925.

the Royal Society of Tropical Medicine and Hygiene 1979;73 483–497.

chondrial DNA sequences. Acta Tropica 2006;99 272–280.

Veterinary Entomology 2007;21 30–35.

Biology 1999;8 381–390.

1994;91 6885–6888.

Parasitology 2002;32 563‒574.

tal Region. Molecular Phylogenetics and Evolution 2009;52 588–601.

2007;101 239–244.

54 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

324 (in Chinese).


## **Systematic Techniques for the Recognition of** *Anopheles* **Species Complexes**

Wej Choochote and Atiporn Saeung

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/54853

## **1. Introduction**

Throughout the world, 528 species of *Anopheles* mosquitoes have been discovered, and approximately 80 of them play an important role as vectors of malaria, filarial nematode and encephalitis virus. Among these, at least 20 taxa represent species complexes, which comprise about 115 sibling species members. The existence of species complexes in *Anopheles* vectors leads to difficulty in precisely identifying sibling species (isomorphic species) and/or subspe‐ cies (morphologically/cytologically polymorphic races) members that possess identical morphology or minimal morphological distinction. In addition, those members may differ in biological characteristics (e.g., microhabitats, resting and biting behavior, sensitivity or resistance to insecticides, susceptible or refractory to malaria parasites, etc.), which can be used to determine their potential for transmitting disease agents. Incorrect identification of indi‐ vidual members in *Anopheles* species complexes may result in failure to distinguish between a vector and non-vector, and lead to complications and/or unsuccessful vector control [1-5].

So far, at least 1 and 2 traditional techniques have been used widely for the recognition of sibling species and/or subspecies members at post- and pre-mating barriers. For post-mating barriers; the hybridization or crossing experiment, using the artificial mating technique to determine hybrid non-viability, sterility or breakdown, is still a useful tool for recognizing *Anopheles* species complexes. Detailed genetic incompatibility, including lack of insemination, embryonation, hatchability, larval survival, pupation, emergence, adult sex distortion, abnormal reproductive system and complete or incomplete (some cases only at the inversion heterozygote regions) asynaptic salivary gland polytene chromosomes are useful criteria for elucidating sibling species and subspecies status. However, a point worth noting is that an isofemale line (isoline) colony established from the combinative characters of morphological and/ or cytological markers has to be considered seriously. A laboratory raised colony established

© 2013 Choochote and Saeung; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Choochote and Saeung; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

from a naturally mixed population should be omitted, since it may be a mixture of cryptic species [6-10]. In addition, many *Anopheles* species do not reproduce in captivity. As for premating barriers; examination of the polytene chromosomes in wild-caught adult females, and/ or progenies of iso-female lines, provides clear evidence that different specific mate recognition systems (SMRS) exist. The total absence or significantly deficient number of heterozygotes for an inversion in a sympatric population entirely indicates the presence of reproductive isolation within a taxon [10-12]. Nonetheless, at least 4 problems have been raised regarding this matter, i.e., (1) a skilled person is needed to prepare a perfect chromosome and make an identification, (2) homosequential banding species cannot be employed, e.g., *An. maculipennis* complex [13] and *An. barbirostris* complex [14-17], (3) a relatively large amount of sample materials are required to perform the Hardy-Weinberg equilibrium, which cannot be applied to small numbers of rare species specimens that are caught during specific seasons, and (4) it cannot be performed in allopatric anopheline populations. Electrophoretic variations at enzyme loci are not only useful for identification of sibling species, but also for the correct identification of morphologically cryptic *Anopheles* species. Variations at a locus thus enable detection of reproductive isolation within populations, resulting from positive assortative (preferential) mating [10-11, 18]. Nevertheless, at least 2 problems have been raised regarding this technique, i.e., (1) specimens must be fresh or frozen until analysis, and (2) its use must be similar to that of the polytene chromosome, as it requires a relatively large amount of sample materials to perform the Hardy-Weinberg equilibrium and cannot be performed in allopatric anopheline populations, as previously described.

Regarding the modernized technique; molecular investigation of some specific genomic markers, e.g., ribosomal DNA (ITS2, D2, D3, IGS) and mitochondrial DNA (COI, COII, Cyt b, ND5), has been used extensively as a tool to characterize and/or diagnose cryptic members in the intra-taxa of *Anopheles* mosquitoes, and the advantage of this PCR-based technique is that few nanograms of DNA are required from preserved specimens [19]. Nonetheless, controversy arose when only comparative DNA sequence analyses of some specific genomic regions were used as first hand criteria to differentiate between the status of specific species, sibling species and subspecies within the taxon *Anopheles*. For example, based on a comparison of the D3 domain of 28S (28S-D3), *An. fluviatilis* S has been considered as synonymous to the *An. minimus* species C [20-22]. However, subsequent investigation of the conspecificity of these two species, based on ITS2 and D2-D3 domains of 28S rDNA regions, suggests that *An. fluviatilis* S and *An. minimus* C, do not deserve to be synonymous [23]. Similar results were also obtained in the determining on specific species status between *An. lesteri* and *An. paraliae* [unpublished data]. The comparative DNA sequence analyses between *An. lesteri* strain from Korea and *An. paraliae* strain from Thailand revealed low pairwise genetic distance for COI (0.007-0.017) and COII (0.008-0.011) regions with 4-9 and 5-7 base substitutions, respectively, whereas a considerable genetic distance (0.040) was obtained in ITS2 region with 16 base substitutions. Supportively, the phylogenetic trees demonstrated that these two species were separated from each other with a 74-100% bootstrap value for 3 regions. It was interesting to note that *An. lesteri* and *An. paraliae* were distinguished appreciably by DNA sequence data, however, were confirmed to be genetically compatible by the crossing experiments. Remark‐ ably, prior to reaching a definite conclusion of specific species, sibling species and subspecies status within the taxon *Anopheles*, crossing experiments need to be carried out intensively using iso-female lines established from sympatric and/or allopatric populations, which relate to morphological variants, cytogenetic forms and/or comparative DNA sequence analyses of some specific regions.

3

populations, which relate to morphological variants, cytogenetic forms and/or comparative

systematic procedures for the recognition of *Anopheles* species complexes [24] (Figure 1).

## **2. Formation of robust systematic procedures**

from a naturally mixed population should be omitted, since it may be a mixture of cryptic species [6-10]. In addition, many *Anopheles* species do not reproduce in captivity. As for premating barriers; examination of the polytene chromosomes in wild-caught adult females, and/ or progenies of iso-female lines, provides clear evidence that different specific mate recognition systems (SMRS) exist. The total absence or significantly deficient number of heterozygotes for an inversion in a sympatric population entirely indicates the presence of reproductive isolation within a taxon [10-12]. Nonetheless, at least 4 problems have been raised regarding this matter, i.e., (1) a skilled person is needed to prepare a perfect chromosome and make an identification, (2) homosequential banding species cannot be employed, e.g., *An. maculipennis* complex [13] and *An. barbirostris* complex [14-17], (3) a relatively large amount of sample materials are required to perform the Hardy-Weinberg equilibrium, which cannot be applied to small numbers of rare species specimens that are caught during specific seasons, and (4) it cannot be performed in allopatric anopheline populations. Electrophoretic variations at enzyme loci are not only useful for identification of sibling species, but also for the correct identification of morphologically cryptic *Anopheles* species. Variations at a locus thus enable detection of reproductive isolation within populations, resulting from positive assortative (preferential) mating [10-11, 18]. Nevertheless, at least 2 problems have been raised regarding this technique, i.e., (1) specimens must be fresh or frozen until analysis, and (2) its use must be similar to that of the polytene chromosome, as it requires a relatively large amount of sample materials to perform the Hardy-Weinberg equilibrium and cannot be performed in allopatric anopheline

Regarding the modernized technique; molecular investigation of some specific genomic markers, e.g., ribosomal DNA (ITS2, D2, D3, IGS) and mitochondrial DNA (COI, COII, Cyt b, ND5), has been used extensively as a tool to characterize and/or diagnose cryptic members in the intra-taxa of *Anopheles* mosquitoes, and the advantage of this PCR-based technique is that few nanograms of DNA are required from preserved specimens [19]. Nonetheless, controversy arose when only comparative DNA sequence analyses of some specific genomic regions were used as first hand criteria to differentiate between the status of specific species, sibling species and subspecies within the taxon *Anopheles*. For example, based on a comparison of the D3 domain of 28S (28S-D3), *An. fluviatilis* S has been considered as synonymous to the *An. minimus* species C [20-22]. However, subsequent investigation of the conspecificity of these two species, based on ITS2 and D2-D3 domains of 28S rDNA regions, suggests that *An. fluviatilis* S and *An. minimus* C, do not deserve to be synonymous [23]. Similar results were also obtained in the determining on specific species status between *An. lesteri* and *An. paraliae* [unpublished data]. The comparative DNA sequence analyses between *An. lesteri* strain from Korea and *An. paraliae* strain from Thailand revealed low pairwise genetic distance for COI (0.007-0.017) and COII (0.008-0.011) regions with 4-9 and 5-7 base substitutions, respectively, whereas a considerable genetic distance (0.040) was obtained in ITS2 region with 16 base substitutions. Supportively, the phylogenetic trees demonstrated that these two species were separated from each other with a 74-100% bootstrap value for 3 regions. It was interesting to note that *An. lesteri* and *An. paraliae* were distinguished appreciably by DNA sequence data, however, were confirmed to be genetically compatible by the crossing experiments. Remark‐ ably, prior to reaching a definite conclusion of specific species, sibling species and subspecies

populations, as previously described.

58 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

In light of the advantages and disadvantages of the techniques mentioned above, 3 techniques, i.e., the crossing experiment, molecular investigation and cytogenetic markers (characteristics of metaphase karyotypes) were selected, and they formed the robust systematic procedures for the recognition of *Anopheles* species complexes [24] (Figure 1). DNA sequence analyses of some specific regions. **Formation of robust systematic procedures**  In light of the advantages and disadvantages of the techniques mentioned above, 3 techniques, i.e., the crossing experiment, molecular investigation and cytogenetic markers (characteristics of metaphase karyotypes) were selected, and they formed the robust

Figure 1. Summarized flow chart for robust systematic procedure **Figure 1.** Summarized flow chart for robust systematic procedures

 By following the flow chart: (1) try to collect anopheline mosquitoes that are distinct in their behavior (e.g., biting humans or animals with relation to different microhabitats and/or locations), (2) try to record morphological variation(s) as far as possible during the species identification process of wild-caught females, (3) establish an iso-female line colony by allowing gravid females to lay eggs individually, (4) conduct molecular investigation of laid-egg feral females to obtained a robust DNA marker, with this step usually taking about 1 week. Since development of the F1-progeny usually takes about 2 weeks from first instar larvae to adults, the metaphase karyotype investigation of fourth instar larvae, newly emerged adult females and males is performed in order to (5) obtain a cytogenetic marker (karyotypic form), (6) if molecular investigation fails in the step of laid-egg feral female it will be performed in F1-progeny, (7) carry out morphometric and morphological investigations of eggs, larvae, pupal skins and adults to confirm precise species identification, and (8) perform the important step of crossing experiments among iso-female line colonies by using a karyotypic marker (or form) related to a DNA marker (large sequence divergence or very low intraspecific sequence variation) of each iso-female line colony. By following the flow chart: (1) try to collect anopheline mosquitoes that are distinct in their behavior (e.g., biting humans or animals with relation to different microhabitats and/or locations), (2) try to record morphological variation(s) as far as possible during the species identification process of wild-caught females, (3) establish an iso-female line colony by allowing gravid females to lay eggs individually, (4) conduct molecular investigation of laidegg feral females to obtained a robust DNA marker, with this step usually taking about 1 week. Since development of the F1-progeny usually takes about 2 weeks from first instar larvae to adults, the metaphase karyotype investigation of fourth instar larvae, newly emerged adult females and males is performed in order to (5) obtain a cytogenetic marker (karyotypic form), (6) if molecular investigation fails in the step of laid-egg feral female it will be performed in F1-progeny, (7) carry out morphometric and morphological investigations of eggs, larvae, pupal skins and adults to confirm precise species identification, and (8) perform the important step of crossing experiments among iso-female line colonies by using a karyotypic marker (or form) related to a DNA marker (large sequence divergence or very low intraspecific sequence variation) of each iso-female line colony.

Regarding techniques necessary for success in operating robust systematic procedure: 3 important techniques were developed by the authors, and they have been proven as efficient and necessary for the robust systematic recognition of sibling species and/or subspecies members within the taxon *Anopheles* species complex. They are: (1) the establishment of a healthy iso-female line colony that is the backbone of population-genetic study on *Anopheles* vectors, since it provides healthy larval and adult progenies for preparation of attractive metaphase and salivary gland polytene chromosomes, and potent adults for crossing experi‐ ments. The inability to establish a healthy iso-female line colony that can be colonized for many consecutive generations is the principle cause of failure in a population-genetic study of *Anopheles* vectors, (2) the technique for metaphase chromosome preparations in adult females and males by intrathoracic inoculation [25] and that for fourth instar larval brains [14] using extracted solution derived from dried seeds and rhizomes of a decoration plant (*Gloriosa superba* L.), instead of synthetic colchicine solution, and (3) modified technique for salivary gland polytene chromosome preparations in fourth instar larvae [26]. Detailed and important procedures regarding the 3 techniques are as follows:

## **3. Techniques for establishment of a healthy iso-female line colony of difficult-to-rear anophelines**

An iso-female line colony of *An. campestris*-like Form E, Thai strain [14] was established from 1 wild-caught fully engorged adult female collected from a human-baited trap reared suc‐ cessfully under laboratory conditions for 98 consecutive generations and used as a role model for other fresh-water breeding anopheline species.

## **4. Procedures**

#### **4.1. Transportation of wild-caught anophelines**

Wild-caught fully engorged adult females collected from human- and/or animal-baited traps in the field were kept in a plastic cup (8.5 cm in diameter and 11 cm in depth, lined inside with filter paper), with a pad of cotton wool soaked with 10% sucrose solution placed on top of the covering screen. It was covered with a translucent plastic bag in order to keep humid condi‐ tions in the cup and delay rapid drying of the soaked cotton wool (Figure 2a). It was stored in a humid chamber using a picnic foam-box (18 x 26 x 39 cm) to maintain humidity and temperature (Figure 2b). Then it was transported to the insectarium for colonization and biological studies. All of the experiments were performed in the insectarium at 27±2 ºC, 70-80% relative humidity, and illumination from a combination of natural daylight from a glasswindow and fluorescent lighting was provided for approximately 12 hours a day.

**Figure 2.** (a) A screen-topped plastic cup with a pad of soaked cotton wool placed on top of the covering screen (left), covered with a translucent plastic bag (right), and a humid chamber derived from a picnic foam-box (background). (b) Top view of the humid chamber showing 6 plastic cups placed on a wet towel lined the bottom (pink colour) and 10-15 ice cubes

#### **4.2. Egg laying**

step of crossing experiments among iso-female line colonies by using a karyotypic marker (or form) related to a DNA marker (large sequence divergence or very low intraspecific sequence

Regarding techniques necessary for success in operating robust systematic procedure: 3 important techniques were developed by the authors, and they have been proven as efficient and necessary for the robust systematic recognition of sibling species and/or subspecies members within the taxon *Anopheles* species complex. They are: (1) the establishment of a healthy iso-female line colony that is the backbone of population-genetic study on *Anopheles* vectors, since it provides healthy larval and adult progenies for preparation of attractive metaphase and salivary gland polytene chromosomes, and potent adults for crossing experi‐ ments. The inability to establish a healthy iso-female line colony that can be colonized for many consecutive generations is the principle cause of failure in a population-genetic study of *Anopheles* vectors, (2) the technique for metaphase chromosome preparations in adult females and males by intrathoracic inoculation [25] and that for fourth instar larval brains [14] using extracted solution derived from dried seeds and rhizomes of a decoration plant (*Gloriosa superba* L.), instead of synthetic colchicine solution, and (3) modified technique for salivary gland polytene chromosome preparations in fourth instar larvae [26]. Detailed and important

**3. Techniques for establishment of a healthy iso-female line colony of**

An iso-female line colony of *An. campestris*-like Form E, Thai strain [14] was established from 1 wild-caught fully engorged adult female collected from a human-baited trap reared suc‐ cessfully under laboratory conditions for 98 consecutive generations and used as a role model

Wild-caught fully engorged adult females collected from human- and/or animal-baited traps in the field were kept in a plastic cup (8.5 cm in diameter and 11 cm in depth, lined inside with filter paper), with a pad of cotton wool soaked with 10% sucrose solution placed on top of the covering screen. It was covered with a translucent plastic bag in order to keep humid condi‐ tions in the cup and delay rapid drying of the soaked cotton wool (Figure 2a). It was stored in a humid chamber using a picnic foam-box (18 x 26 x 39 cm) to maintain humidity and temperature (Figure 2b). Then it was transported to the insectarium for colonization and biological studies. All of the experiments were performed in the insectarium at 27±2 ºC, 70-80% relative humidity, and illumination from a combination of natural daylight from a glass-

window and fluorescent lighting was provided for approximately 12 hours a day.

variation) of each iso-female line colony.

60 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

procedures regarding the 3 techniques are as follows:

for other fresh-water breeding anopheline species.

**4.1. Transportation of wild-caught anophelines**

**difficult-to-rear anophelines**

**4. Procedures**

After the engorged adult female was maintained for 4-5 days and/or until gravid in the insectarium, it was placed in a screen-topped oviposition plastic-cup (6 cm in diameter and 7 cm in depth) containing 25 ml of natural water (brought from a basin that was used for tapwater production). Wet filter paper lined the inside of the screen-topped was covered with a black plastic sheet (Figure 3a-c). The eggs attached to the moist side of the filter paper and/or floating on the water surface were rinsed and transferred to white plastic tray (25 x 36 x 6 cm) containing 1,500 ml rearing water (equal part of natural water and distilled water) with wet filter paper lining the inside. During the embryonation period, the eggs were exposed to a 40 watt light instead of sunlight, for warming the eggs until hatching (Figure 3d).

#### **4.3. Rearing of larvae, pupae and adults**

After egg hatching, first instar larvae were transferred daily from an ovipot to a white plastic tray (25 x 36 x 6 cm) containing 2,000 ml rearing water and approximately 15 stems of garden grass (*Axonopus compressus*), and 80 first instar larvae were reared in each tray. The rearing tray was covered with a transparent plastic sheet for reducing the need to change and/or refill the tray with rearing water during the larval development process (Figure 4a-b). An extra and/or a standard formula of fish food consisting approximately of protein 47.5%, oil 6.5%, fibre 2.0%, ash 10.5%, moister 6.0% and additives of vitamins A (29,770 IU/kg), D3 (1,860 IU/ kg), E (200 mg/kg), L-ascorbyl-2-polyphosphate (138 mg/kg), lecithin, l-lysine monochlorhy‐ drate, and citric acid was used as larval nutrient. Fine fish food was placed in a vial covered with a nylon screen (34 x 43 threads per cm2 ) and sprinkled on the water until the food particles stopped spreading across the water's surface. First and second instar larvae were fed twice daily, and this schedule was increased to 3-5 times daily after most of the larvae reached third and fourth instars, respectively. Before each feeding, floating clumps of excess food were

**Figure 3.** (a) A screen-topped oviposition plastic-cup, (b) covered with a black plastic sheet, and (c) top view of the plastic cup showing egg-batch after 12-hours-oviposition of a gravid adult female. (d) Eggs placed in a white plastic tray and exposed to a 40-watt light

removed by dragging a sheet of typing-paper across the water's surface. Any larvae trapped on the paper during the cleaning process were dislodged by rinsing the paper in a tray of rearing water and returning it to the rearing tray. After pupation, approximately 100 pupae placed in a plastic cup (14.5 cm in diameter and 6 cm in depth) containing 150 ml of distilled water were kept in a 30 x 30 x 30 cm cage, and the emerged adults were provided with both 10% sucrose solution and 5% multivitamin syrup solution (consisting approximately of vitamins A: 2,000 I.U., D: 200 I.U., E: 1.50 I.U., B1: 0.70 mg, B2: 0.85 mg, B6: 0.35 mg and C: 17.50 mg, nicotinamide: 9.00 mg, orange juice: 0.50 g and cod liver: 0.10 g per 100 ml solution) saturated in cotton wool coiled around a small piece of wood and placed in a small bottle. Increased humidity to promote adult survival was provided by covering the cage with a wet towel overlaid with a black plastic sheet (Figure 4c). One-day-old males were removed daily from the cage and kept in a screen-topped plastic cup (lined inside with filter paper), where they were provided with a 5% multivitamin syrup solution through a pad of soaked cotton wool, which was placed on top of the screen and changed daily. In order to keep humid conditions in the cup and delay rapid drying of the cotton wool soaked in 5% multivitamin syrup solution, the screen-top was covered with a translucent plastic bag (Figure 4d).

**Figure 4.** (a) Top view of a white plastic tray placed with 15 stems of garden grass, and (b) covered with a transparent plastic sheet. (c) Adult rearing cage partially covered with a wet towel (pink colour) and a black plastic-sheet with plas‐ tic container for holding pupae, and two bottles with cotton wicks, one containing 10% sucrose solution and another 5% multivitamin syrup solution. (d) Adult males being kept in a screen-topped plastic cup (lined inside with filter pa‐ per) with a pad of cotton wool soaked in 5% multivitamin syrup solution and the top covered with a translucent plas‐ tic bag to maintain humidity.

#### **4.4. Suitable blood-feeding condition**

removed by dragging a sheet of typing-paper across the water's surface. Any larvae trapped on the paper during the cleaning process were dislodged by rinsing the paper in a tray of rearing water and returning it to the rearing tray. After pupation, approximately 100 pupae placed in a plastic cup (14.5 cm in diameter and 6 cm in depth) containing 150 ml of distilled water were kept in a 30 x 30 x 30 cm cage, and the emerged adults were provided with both 10% sucrose solution and 5% multivitamin syrup solution (consisting approximately of vitamins A: 2,000 I.U., D: 200 I.U., E: 1.50 I.U., B1: 0.70 mg, B2: 0.85 mg, B6: 0.35 mg and C: 17.50 mg, nicotinamide: 9.00 mg, orange juice: 0.50 g and cod liver: 0.10 g per 100 ml solution) saturated in cotton wool coiled around a small piece of wood and placed in a small bottle. Increased humidity to promote adult survival was provided by covering the cage with a wet towel overlaid with a black plastic sheet (Figure 4c). One-day-old males were removed daily from the cage and kept in a screen-topped plastic cup (lined inside with filter paper), where they were provided with a 5% multivitamin syrup solution through a pad of soaked cotton wool, which was placed on top of the screen and changed daily. In order to keep humid conditions in the cup and delay rapid drying of the cotton wool soaked in 5% multivitamin

**Figure 3.** (a) A screen-topped oviposition plastic-cup, (b) covered with a black plastic sheet, and (c) top view of the plastic cup showing egg-batch after 12-hours-oviposition of a gravid adult female. (d) Eggs placed in a white plastic

tray and exposed to a 40-watt light

62 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

syrup solution, the screen-top was covered with a translucent plastic bag (Figure 4d).

Comparative direct feeding ability on white rat in a 30 x 30 x 30 cm cage, and artificial feeding ability on human heparinized-blood (obtained from human volunteers whom sign the consent form) in a plastic cup (8.5 cm in diameter and 11 cm in depth, lined inside with filter paper) (Figure 5), of female *An. campestris*-like Form E at different ages ranging from 1 to 10 days, demonstrated that in the cage, adult females aged of 3, 4, 5 and 6 days were successful in feeding on the blood of white rats, with feeding rates of 30%, 39%, 62% and 43%, respectively. Interestingly, the adult females aged 3, 4, 5, and 6 days succeeded in artificial feeding on human heparinized-blood in the plastic cup at higher rates than direct feeding on white rat in the cage in all experiments by yielding feeding rates of 62%, 68%, 78% and 61%, respectively. Never‐ theless, the engorged females that derived from 2 feeding methods were used satisfactorily for the maintenance of an iso-female line laboratory-raised colony of *An*. *campestris*-like Form E. One difficulty and/or failure in rearing mosquitoes in the laboratory was the subsequent generation's refusal to feed on blood, particularly from small laboratory animals such as guinea pig, white rat, golden hamster, etc. This leads to direct feeding from human volunteers, especially at the beginning of the first to fifth generations of the colony. However, to solve this problem, forced artificial feeding on human heparinized-blood by *An. campestris*-like Form E was successful in this study and has been used routinely up to this time. Nonetheless, a point to be kept in mind is that only the healthy progenies of laboratory-raised colonies could be used successfully. Additionally, the use of direct blood feeding of subsequent mosquito progenies from human volunteers is a potentially dangerous method and should be given up entirely, since at least 4 reports have declared that *An. peditaeniatus* [27], *An. subpictus* [28-29] and *An. barbirostris* [30] have been incriminated as secondary vectors of Japanese encephalitis virus, which is possibly transmitted vertically.

**Figure 5.** Artificial feeding system. A warm water-bath at 40ºC, with a water pump placed inside, is connected to glass inlet and outlet feeding-chambers by rubber tubes. Thin paraffin-membrane covers the bottom tip of the feeding chambers, which are filled with human heparinized-blood, and the bottom tip is in close contact with 50 fasted adult female *An. campestris*-like Form E that are inside a screen-topped paper cup.

#### **4.5. Ability of free mating in a 30 cm cubed cage and male ability to mate artificially**

One of the difficulties in the colonization of anopheline mosquitoes in the laboratory might be due to adults not being capable of copulation in a small and/or standard cage (30 x 30 x 30 cm). Thus, in order to determine the adaptive stenogamy of *An. campestris*-like Form E, the newly emerged females and males co-habitated at a ratio of 200/300, in a 30 x 30 x 30 cm cage for one week [31-32]. The results indicated that *An*. *campestris*-like Form E failed to mate freely in the cage at a 0% insemination rate (from experiments repeated 3 times), indicating strong euryg‐ amy. Thus, the artificial mating methods as described by [33-34] were used. The best age for artificial mating in male *An. campestris-*like Form E was 5-days-old (100% mating rate, 86.67% insemination rate). Nonetheless, males aged 4 and 8 days old could be used satisfactorily (93.33-100% mating rates, 80-82.14% insemination rates) (Table 1).


**Table 1.** Artificial mating ability of *An. campestris*-like Form E males

generation's refusal to feed on blood, particularly from small laboratory animals such as guinea pig, white rat, golden hamster, etc. This leads to direct feeding from human volunteers, especially at the beginning of the first to fifth generations of the colony. However, to solve this problem, forced artificial feeding on human heparinized-blood by *An. campestris*-like Form E was successful in this study and has been used routinely up to this time. Nonetheless, a point to be kept in mind is that only the healthy progenies of laboratory-raised colonies could be used successfully. Additionally, the use of direct blood feeding of subsequent mosquito progenies from human volunteers is a potentially dangerous method and should be given up entirely, since at least 4 reports have declared that *An. peditaeniatus* [27], *An. subpictus* [28-29] and *An. barbirostris* [30] have been incriminated as secondary vectors of Japanese encephalitis

**Figure 5.** Artificial feeding system. A warm water-bath at 40ºC, with a water pump placed inside, is connected to glass inlet and outlet feeding-chambers by rubber tubes. Thin paraffin-membrane covers the bottom tip of the feeding chambers, which are filled with human heparinized-blood, and the bottom tip is in close contact with 50 fasted adult

One of the difficulties in the colonization of anopheline mosquitoes in the laboratory might be due to adults not being capable of copulation in a small and/or standard cage (30 x 30 x 30 cm). Thus, in order to determine the adaptive stenogamy of *An. campestris*-like Form E, the newly emerged females and males co-habitated at a ratio of 200/300, in a 30 x 30 x 30 cm cage for one week [31-32]. The results indicated that *An*. *campestris*-like Form E failed to mate freely in the cage at a 0% insemination rate (from experiments repeated 3 times), indicating strong euryg‐ amy. Thus, the artificial mating methods as described by [33-34] were used. The best age for artificial mating in male *An. campestris-*like Form E was 5-days-old (100% mating rate, 86.67% insemination rate). Nonetheless, males aged 4 and 8 days old could be used satisfactorily

**4.5. Ability of free mating in a 30 cm cubed cage and male ability to mate artificially**

female *An. campestris*-like Form E that are inside a screen-topped paper cup.

(93.33-100% mating rates, 80-82.14% insemination rates) (Table 1).

virus, which is possibly transmitted vertically.

64 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

#### **4.6. Searching for a suitable oviposition-condition**

Many anopheline colonies have been reported to adapt easily to oviposit eggs in the cage on various types of simple ovipots, e.g., petridish, crystallizing dish, terra-cotta bowl, white plastic cup, black cup, etc. [35-39]. In the case of using 20 gravid adult females of *An. campest‐ ris*-like Form E put in a 30 x 30 x 30 cm cage for 12 hours (starting from 18.00-06.00 hours), the results revealed that 0, 0, 279, 0 and 0 eggs per an oviposited-plastic cup (9 cm in diameter and 10.5 cm in depth, containing 80 ml of natural water) were found in experiments 1, 2, 3, 4 and 5, respectively; whereas the forced laying of eggs by placing 20 gravid adult females in an oviposited-plastic cup (details mentioned above in paragraph 2 "Egg laying") in the same size and conditions as used in the cage, a massive number of eggs, i.e., 1,273, 1,318, 1,705, 2,180 and 1,501 eggs per cup, were recovered for experiments 1, 2, 3, 4 and 5, respectively (Figure 6). The high yield of eggs recovered from the latter experiment appears to result in the fact that the close-system of an oviposited-plastic cup provided significantly higher relative humidity than a cage or open-system. The air-rich water molecules in high relative humidity are the important attractants to gravid female alfactometer, which indicates suitable or acceptable oviposition sites [40]. Thus, in oviposition of *An. campestris*-like Form E and other anopheline species in our laboratory, this method has been used routinely up until now.

**Figure 6.** Top view of ovipot derived from a plastic cup showing massive egg-batches after 12-hours-oviposition of the 20 gravid adult females

#### **4.7. Other important factors**

Throughout the larval rearing period, the number of larvae, rearing conditions in the tray, and food were the most important factors, not only for routine rearing, but also special rearing in order to obtain a high yield of metaphase and polytene chromosomes, which were necessary for population-genetic study of anophelines. Stressful rearing-conditions, e.g., the overcrowd‐ ing of larvae in a rearing tray (in this study, 80 larvae per 25 x 36 x 6 cm tray was an appropriate number for *An. campestris*-like Form E), and the use of inappropriate water medium and food would lead to a rapid drop in and/or loss of a colony. Also, this would result in low larval and pupal survival rates, adult F1-progenies refusing to take blood meal, difficulty in artificial mating of adult females and males and/or failure to inseminate sperm into mated-female spermathecae, short life span of adult females and males, mated gravid adult females laying fewer numbers of eggs and/or failure to lay eggs, and low egg-hatchability. Thus, any rearing system, which is an important first step that leads to obtaining healthy larvae, would be a promising method for successfully establishing a colony, particularly an iso-female line colony, which is more difficult and complicated to establish than a mixed colony. As mentioned previously, food was one of the most important factors for obtaining healthy larvae, thus, several kinds of larval food were tested for use and comparison, e.g., mouse pellets, cat and dog biscuits and various formulas of fish food. The results indicated that the standard formula of fish food as mentioned in paragraph 3 ("Rearing of larvae, pupae and adults"), proved to be an excellent larval food for *An. campestris-*like Form E. It is expected that this fish food formula was also ideal for other anopheline species with rearing difficulties. The use of equal part of natural water and distilled water as the larval rearing medium also proved to be promising. Trials using boiled tap-water, filtered tap-water, polarized water and deionized water yielded unsatisfactory outcomes by providing low larval survival, particularly through subsequent progenies. The addition of garden grass to the larval rearing tray, as stated by [31], resulted in high larval survival for *An. campestris-*like Form E. Using few stems of garden grass, or withdrawing it, would lead to low larval survival and/or weak larvae for rearing subsequent generations. Using slightly more or less than 15 stems of garden grass, depending upon the size of the stems, and size and number of leaves, proved to improve conditions to a suitable level for larval rearing, since the grass provided a resting place for larvae, rendered shade as in natural breeding sites (rice paddy, ponds and swamps associated with water plants) [41-42], and aerated the medium. Its roots were also very important for maintaining clear and clean rearing medium by using larval waste products and unconsumed food as fertilizer, which determined the obvious active growth of grass in the rearing tray. Finally, we hope that the detailed information concerning rearing aspects of *An. campestris*-like Form E will prove to be important for the establishment of other anopheline species that have been previously difficult to rear.

**Notes**: By following the systematic rearing procedures as detail-mentioned above, at least 23 *Anopheles* species were successful reared in our insectarium, i.e., subgenus *Anopheles* [*An. argyropus*(F23), *An. barbirostris* species A1 (F86), *An. belenrae*(F26), *An. campestris*-like Form E (F98), *An. crawfordi* (F23), *An. lesteri* (F60), *An. nigerrimus* (F23), *An. nitidus* (F28), *An. paraliae* (F24), *An. peditaeniatus* (F23), *An. pullus* (F24), *An. pursati* (F24) and *An. sinensis* (F28)]; and *Cellia* [*An. harrisoni* (F51), *An. jamesii* (F10), *An*. *jeyporiensis* (F5), *An. karwari* (F13), *An. kochi* (F25), *An. nivipes* (F12), *An. pampanai* (F11), *An. philippinensis* (F12), *An. splendidus* (F10) and *An. tessellatus* (F27)].

## **5. Techniques for metaphase and polytene chromosome preparations**

#### **5.1. Rearing condition of mosquitoes for chromosome preparations**

**Figure 6.** Top view of ovipot derived from a plastic cup showing massive egg-batches after 12-hours-oviposition of

Throughout the larval rearing period, the number of larvae, rearing conditions in the tray, and food were the most important factors, not only for routine rearing, but also special rearing in order to obtain a high yield of metaphase and polytene chromosomes, which were necessary for population-genetic study of anophelines. Stressful rearing-conditions, e.g., the overcrowd‐ ing of larvae in a rearing tray (in this study, 80 larvae per 25 x 36 x 6 cm tray was an appropriate number for *An. campestris*-like Form E), and the use of inappropriate water medium and food would lead to a rapid drop in and/or loss of a colony. Also, this would result in low larval and pupal survival rates, adult F1-progenies refusing to take blood meal, difficulty in artificial mating of adult females and males and/or failure to inseminate sperm into mated-female spermathecae, short life span of adult females and males, mated gravid adult females laying fewer numbers of eggs and/or failure to lay eggs, and low egg-hatchability. Thus, any rearing system, which is an important first step that leads to obtaining healthy larvae, would be a promising method for successfully establishing a colony, particularly an iso-female line colony, which is more difficult and complicated to establish than a mixed colony. As mentioned previously, food was one of the most important factors for obtaining healthy larvae, thus, several kinds of larval food were tested for use and comparison, e.g., mouse pellets, cat and dog biscuits and various formulas of fish food. The results indicated that the standard formula of fish food as mentioned in paragraph 3 ("Rearing of larvae, pupae and adults"), proved to be an excellent larval food for *An. campestris-*like Form E. It is expected that this fish food formula was also ideal for other anopheline species with rearing difficulties. The use of equal part of natural water and distilled water as the larval rearing medium also proved to be promising. Trials using boiled tap-water, filtered tap-water, polarized water and deionized water yielded unsatisfactory outcomes by providing low larval survival, particularly through subsequent progenies. The addition of garden grass to the larval rearing tray, as stated by [31], resulted in high larval survival for *An. campestris-*like Form E. Using few stems of garden grass,

the 20 gravid adult females

**4.7. Other important factors**

66 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

The methods for rearing conditions were generally routine as mentioned in paragraph 3, except, 10 first instar larvae per tray were used to obtain a high yield of metaphase chromo‐ somes from larval brains, ovaries and testes, and polytene chromosomes from larval salivary glands. Comparative outcome rates of metaphase chromosomes from larval brains and polytene chromosomes from larval salivary glands between routine (80 larvae) and special (10 larvae) rearing revealed as follows: (1) metaphase chromosomes: experiment 1 [10 larvae (87.50%) vs. 80 larvae (33.33%)], 2 [10 larvae (75.00%) vs. 80 larvae (30.00%)] and 3 [10 larvae (77.78%) vs. 80 larvae (30.00%)]; and (2) salivary gland polytene chromosomes: experiment 1 [10 larvae (80.00%) vs. 80 larvae (50.00%)], 2 [10 larvae (66.67%) vs. 80 larvae (50.00%)] and 3 [10 larvae (100.00%) vs. 80 larvae (66.67%)]. Thus, a special rearing with 10 larvae was used routinely for chromosome preparation.

#### **5.2. Preparation of metaphase chromosomes from adult females and males and fourth instar larvae**

#### *5.2.1. Preparation of 0.5% and 1% solutions of dried* Gloriosa superba *seed and rhizome powders*

Summarized flow chart for normal saline-extracted *Gl. superba* seed and rhizome powders, as follows:

\*By keeping at this condition, the colchicine-like activity in the filtrate stays stable for at least 2 years.

**Notes:** colchicine solution has been used widely at a concentration of 0.05-1% for metaphase chromosome preparation in the cytogenetic study of eukaryotic organisms, e.g., protozoans [43], helminthes [44-45], snails [46], insects [8, 47-50], and plants [51-52]. Spindle formation or microtubule polymerization inhibits arresting mitosis at the metaphase [53-54]. The alkaloid colchicine was isolated from a plant named autumn crocus or meadow saffron (*Colchicum autumnale* L., Family Liliaceae) in 1820 by Pelletier and Caventou [53]. At present, the com‐ mercial products derived from this plant are merchandised extensively and used worldwide. Recently, systematic and continuous studies evaluated the colchicine-like activity of a common decorative plant found widely in tropical countries, Dong Deung (*Gl. superba*, Family Liliaceae) [55], which highlighted the benefits of this plant used for metaphase chromosome preparation in mosquitoes [14, 24, 56-58]. Various concentrations and/or extracted-fractions of dried *Gl. superba* seed and rhizome powders yielded similar metaphase rates and an average number of metaphase chromosomes per positive mosquito to synthetic colchicine solution, indicating that these extracts could be used to replace colchicine. In addition, the authors also mentioned that considerable budget savings could be realized by using their techniques.

Other benefits include a decorative plant that can be bought at many shops in Thailand's flower-markets, and it is hoped elsewhere in tropical countries. It can be grown easily in smallspaced land and outdoors with general fertilizers (e.g., simple formula chemical fertilizer, organic fertilizer and animal manure), which are necessary to promote its growth. It takes about 5-7 months to grow from small budding-rhizomes into mature tree with flowers and green pods (Figure 7a-d).

**Figure 7.** Showing a common decorative plant, Dong Deung (*Gl. superba*). (a) Dong Deung trees with beautiful flow‐ ers and green pods, (b) Ripe and broken Dong Deung pods with reddish-orange seeds, (c) Dried Dong Deung seeds and (d) Dried Dong Deung rhizomes

*5.2.2. Preparation of the metaphase chromosomes from adult females and males and fourth instar larvae*

#### *5.2.2.1. Procedures*

**Notes:** colchicine solution has been used widely at a concentration of 0.05-1% for metaphase chromosome preparation in the cytogenetic study of eukaryotic organisms, e.g., protozoans [43], helminthes [44-45], snails [46], insects [8, 47-50], and plants [51-52]. Spindle formation or microtubule polymerization inhibits arresting mitosis at the metaphase [53-54]. The alkaloid colchicine was isolated from a plant named autumn crocus or meadow saffron (*Colchicum autumnale* L., Family Liliaceae) in 1820 by Pelletier and Caventou [53]. At present, the com‐ mercial products derived from this plant are merchandised extensively and used worldwide. Recently, systematic and continuous studies evaluated the colchicine-like activity of a common decorative plant found widely in tropical countries, Dong Deung (*Gl. superba*, Family Liliaceae) [55], which highlighted the benefits of this plant used for metaphase chromosome preparation in mosquitoes [14, 24, 56-58]. Various concentrations and/or extracted-fractions of dried *Gl. superba* seed and rhizome powders yielded similar metaphase rates and an average number of metaphase chromosomes per positive mosquito to synthetic colchicine solution, indicating that these extracts could be used to replace colchicine. In addition, the authors also mentioned

\*By keeping at this condition, the colchicine-like activity in the filtrate stays stable for at least 2 years.

Filtrated 1 ml of filtrate

filled in a 1.5-ml microcentrifuge tube, and kept at -20 C\*

Macerated in 100 ml of 0.85% sodium chloride solution for 6 hours at 27±2 C, and kept at 4 C for 14-16 hours (or in a refrigerator)

that considerable budget savings could be realized by using their techniques.

green pods (Figure 7a-d).

0.5 g of dried seed powder or 1 g of dried rhizome powder

68 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Other benefits include a decorative plant that can be bought at many shops in Thailand's flower-markets, and it is hoped elsewhere in tropical countries. It can be grown easily in smallspaced land and outdoors with general fertilizers (e.g., simple formula chemical fertilizer, organic fertilizer and animal manure), which are necessary to promote its growth. It takes about 5-7 months to grow from small budding-rhizomes into mature tree with flowers and

Metaphase chromosomes for adult females and males were prepared using the modified techniques described by [25]. The newly emerged adult females and males aged up to about 6-12 hours were starved, anaesthetized with ether and placed on their side on a slide under a binocular microscope. A needle was made by drawing out a glass capillary tube in a flame until the pointed end was approximately 80-100 µm in diameter; the shorter the needle the easier it was to handle. An inoculation was made into the post-spiracular area of the meso‐ thorax, and a filtrate of 0.5% solution of dried *Gl. superba* seed powder was introduced into each mosquito by gently blowing down the attached rubber tube. The volume of inoculums could be controlled by observing the extension of abdomen until it was similar in size to the fully-engorged mosquitoes post fed on 10% sucrose solution. A few minutes after inoculation, most of the mosquitoes had recovered completely. Five inoculated mosquitoes were then kept in a 10-ml test tube (1.5 cm in diameter and 10 cm in length), with cotton wool soaked by 3 drops of distilled water closing the opened-side in order to provide adequate moisture. Then, the cotton wool was sealed with paraffin and the test tube held in an insectarium at 27±2 °C and 70-80% relative humidity for 3 hours (Figure 8a-c).

**Figure 8.** (a) Lower row: 1 ml filtrate of 0.5% solution of dried *Gl. superba* seed powder filled in a 1.5-ml microcentri‐ fuge tube, and upper row: an inoculation glass-needle filled with a filtrate. (b) Intra-thoracic inoculation of a filtrate into the post-spiracular area of the mesothorax. (c) Five inoculated mosquitoes kept in a 10-ml test tube

The inoculated mosquitoes were dissected in a small drop of 1% hypotonic sodium citrate solution on a siliconized slide by pulling out the last abdominal segment to obtain the ovaries or testes under a binocular microscope. The organs obtained were left in 1% hypotonic sodium citrate solution for 10 minutes, and then transferred to a small drop of Carnoy's fixative on a siliconized slide for at least 2 minutes. Then, a drop of 60% acetic acid was added, and the organs were torn and mixed well with dissecting needles. A drop of cell suspension was placed on a clean microscopic slide on a warming plate at about 45–50°C. Droplets of cells were released slowly from a Pasteur pipette to form a circular trail of monolayer cells. The dried slides were stained with 20% Giemsa in phosphate buffer pH 7.2 for 1 hour, rinsed with deionized water, air-dried at room temperature, mounted in Permount® (Fisher, Fairlawn, NJ, USA) and examined under a green filter compound microscope. Metaphase karyotypes were identified by following the standard descriptions (Figure 9) [59-60].

could be controlled by observing the extension of abdomen until it was similar in size to the fully-engorged mosquitoes post fed on 10% sucrose solution. A few minutes after inoculation, most of the mosquitoes had recovered completely. Five inoculated mosquitoes were then kept in a 10-ml test tube (1.5 cm in diameter and 10 cm in length), with cotton wool soaked by 3 drops of distilled water closing the opened-side in order to provide adequate moisture. Then, the cotton wool was sealed with paraffin and the test tube held in an insectarium at 27±2 °C

**Figure 8.** (a) Lower row: 1 ml filtrate of 0.5% solution of dried *Gl. superba* seed powder filled in a 1.5-ml microcentri‐ fuge tube, and upper row: an inoculation glass-needle filled with a filtrate. (b) Intra-thoracic inoculation of a filtrate

The inoculated mosquitoes were dissected in a small drop of 1% hypotonic sodium citrate solution on a siliconized slide by pulling out the last abdominal segment to obtain the ovaries or testes under a binocular microscope. The organs obtained were left in 1% hypotonic sodium citrate solution for 10 minutes, and then transferred to a small drop of Carnoy's fixative on a siliconized slide for at least 2 minutes. Then, a drop of 60% acetic acid was added, and the organs were torn and mixed well with dissecting needles. A drop of cell suspension was placed on a clean microscopic slide on a warming plate at about 45–50°C. Droplets of cells were released slowly from a Pasteur pipette to form a circular trail of monolayer cells. The dried slides were stained with 20% Giemsa in phosphate buffer pH 7.2 for 1 hour, rinsed with deionized water, air-dried at room temperature, mounted in Permount® (Fisher, Fairlawn, NJ, USA) and examined under a green filter compound microscope. Metaphase karyotypes were

into the post-spiracular area of the mesothorax. (c) Five inoculated mosquitoes kept in a 10-ml test tube

identified by following the standard descriptions (Figure 9) [59-60].

and 70-80% relative humidity for 3 hours (Figure 8a-c).

70 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

**Figure 9.** Metaphase chromosomes of *An. paraliae* Form A. (a) Ovary chromosomes, showing homozygous large sub‐ metacentric X3 chromosomes. (b) Testis chromosomes, showing large submetacentric X3 and small telocentric Y1 chro‐ mosomes

The techniques for metaphase chromosome preparations in fourth instar larvae mainly followed those described above, except for the 5 fourth instar larvae that were incubated with a 1 ml filtrate of 0.5% dried *Gl. superba* seed powder solution in a 10-ml test tube for two hours. Then, the larval brains were excised, fixed, smeared, stained with Giemsa, mounted and examined under a green filter compound microscope (Figure 10).

**Figure 10.** Metaphase chromosomes from brains of *An. campestris-*like Form E. (a) Showing homozygous submeta‐ centric X2 chromosomes. (b) Showing submetacentric X2 and small metacentric Y5 chromosomes

#### **5.3. Preparation of the polytene chromosome from larval salivary glands**

#### *5.3.1. Procedures*

Salivary gland polytene chromosomes were prepared using the slightly modified published techniques [26, 61]. The early fourth instar larvae were removed from the rearing tray by a dropper and rinsed in clean distilled water. A healthy larva with flared-thorax in appearance was picked up with forceps, attached to filter paper to remove excess water, placed on a siliconized slide filled with a drop of 1% hypotonic sodium citrate solution, and then dissected under a binocular microscope. The head was cut off, and one dissecting needle was inserted through the anterior end of thorax to posterior end. Then, another dissecting needle was scratched along the line of the inserted needle to tear the thorax integument, open the thorax and take out the internal organs before the thorax and abdomen were transferred into a drop of 15% acetic acid on a siliconized slide. The bilobed salivary glands were removed from the thorax using dissecting needles, and only the whitish anterior lobe of each salivary gland was transferred into a small drop of 45% acetic acid on a siliconized slide and left for 1 minute. After that, one drop of 2% aceto-lactic orcein stain was added. After 15 minutes of staining, a grease-free 22 mm2 coverslip was placed on the stained salivary glands. The preparation was wrapped firmly in filter paper and gently pressed with a thumb to squash and spread the chromosomes. Then, the coverslip edges were sealed with transparent nail varnish. The prepared chromosomes were scrutinized under a green filter compound microscope. The arm of the polytene chromosomes was identified by following the standard map (Figure 11) [61].

**Figure 11.** (a) Complete synaptic salivary gland polytene chromosome of *An.campestris*-like Form E. (b) Homosequen‐ tial asynapsis in all autosomes and the X chromosome from crosses between *An.campestris*-like Form E and An. barbir‐ *ostris* species A1

**Notes:** by application of this robust systematic procedure, 5 sibling species members have recently been recognized in the taxon *An. barbirostris* complex within 2 years [14-16]. In addition, 8 species comprising a total of 26 subspecies (cytological forms) have been recognized during the past decade, i.e*., An. vagus* Forms A and B [62], *An. pullus* Forms A and B (= *An*. *yatsushiroensis*) [62], *An. sinensis* Forms A and B [64-66], *An. aconitus* Forms B and C [67], *An. barbirostris* species A1 (Forms A, B, C and D) and A2 (Forms A and B) [14-16], *An. campestris*like Forms B, E, and F [68], *An. peditaeniatus* Forms B, C, D, E [69], and *An. paraliae* Forms A, B, C, D and E [unpublished data].

## **6. Conclusion**

**5.3. Preparation of the polytene chromosome from larval salivary glands**

Salivary gland polytene chromosomes were prepared using the slightly modified published techniques [26, 61]. The early fourth instar larvae were removed from the rearing tray by a dropper and rinsed in clean distilled water. A healthy larva with flared-thorax in appearance was picked up with forceps, attached to filter paper to remove excess water, placed on a siliconized slide filled with a drop of 1% hypotonic sodium citrate solution, and then dissected under a binocular microscope. The head was cut off, and one dissecting needle was inserted through the anterior end of thorax to posterior end. Then, another dissecting needle was scratched along the line of the inserted needle to tear the thorax integument, open the thorax and take out the internal organs before the thorax and abdomen were transferred into a drop of 15% acetic acid on a siliconized slide. The bilobed salivary glands were removed from the thorax using dissecting needles, and only the whitish anterior lobe of each salivary gland was transferred into a small drop of 45% acetic acid on a siliconized slide and left for 1 minute. After that, one drop of 2% aceto-lactic orcein stain was added. After 15 minutes of staining, a grease-free 22 mm2 coverslip was placed on the stained salivary glands. The preparation was wrapped firmly in filter paper and gently pressed with a thumb to squash and spread the chromosomes. Then, the coverslip edges were sealed with transparent nail varnish. The prepared chromosomes were scrutinized under a green filter compound microscope. The arm of the polytene chromosomes was identified by following the standard map (Figure 11) [61].

**Figure 11.** (a) Complete synaptic salivary gland polytene chromosome of *An.campestris*-like Form E. (b) Homosequen‐ tial asynapsis in all autosomes and the X chromosome from crosses between *An.campestris*-like Form E and An. barbir‐

**Notes:** by application of this robust systematic procedure, 5 sibling species members have recently been recognized in the taxon *An. barbirostris* complex within 2 years [14-16]. In addition, 8 species comprising a total of 26 subspecies (cytological forms) have been recognized during the past decade, i.e*., An. vagus* Forms A and B [62], *An. pullus* Forms A and B (= *An*.

*5.3.1. Procedures*

72 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

*ostris* species A1

The formation of robust systematic procedures is highly anticipated, based on the crossing experiments between iso-female lines using cytological markers (characteristics of metaphase chromosomes/karyotypic forms). Together with this information, the data on comparative sequence analyses of some specific genomic regions (rDNA and mtDNA) would bring success in recognizing and reliably identifying sibling species and/or subspecies members within the taxon of other *Anopheles* species complexes. In addition, the detailed techniques necessary for the establishment of difficult-to-rear anopheline species, which yield high rates of attractive metaphase and polytene chromosomes and potent adults for crossing experiments, would be main keys leading to successful study on the population-genetic structure of *Anopheles* vectors. These factors are important for studying the biology, behavior of *Anopheles* species, as well as for an epidemiology and a control approach of the targeted vector species.

## **Acknowledgements**

Sincere thanks are extended to the Thailand Research Fund (TRF Advanced Research Scholar: BRG/14/2545 and BRG5380021, and TRF Senior Research Scholar: RTA5480006), the Thailand Research Fund through the Royal Golden Jubilee Ph.D. Program (Grant No. PHD/0044/2546, PHD/0052/2548, PHD/0082/2549, PHD/0031/2550, PHD/0297/2551 and PHD/0356/2552), Biodiversity Research and Training Program (Grant No. BRT R\_249004, 250009 and 252005: 2006-2009), and Faculty of Medicine Research Fund, Chiang Mai University, Chiang Mai, Thailand, for their continuous financial support in the population-genetic study of *Anopheles* vectors in Thailand.

## **Author details**

Wej Choochote\* and Atiporn Saeung

\*Address all correspondence to: wchoocho@mail.med.cmu.ac.th

Department of Parasitology, Faculty of Medicine, Chiang Mai University, Chiang Mai, Thailand

### **References**


[13] Coluzzi M (1970) Sibling species in *Anopheles* and their importance to malariology. Misc. Publ. Ent. Soc. Am. 7: 63-77.

**References**

46-48.

South-East Asia, New Delhi. 102 p.

74 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Jap. J. Sanit. Zool. 32: 321-329.

Wash. 89: 157-166.

Control Assoc. 6: 477-481.

Publ. SEARO. 18: 1-82.

Vet. Entomol. 6: 335-341.

330-331.

in Assam (India). Acta Trop. 113: 241-244.

Culicidae) in Thailand. Genome 30: 372–379.

[1] Foley DH, Beebe N, Torres E, Saul A (1996) Misidentification of a Philippine malaria vector revealed by allozyme and ribosomal DNA markers Am. J. Trop. Med. Hyg. 54:

[2] Van Bortel W, Harbach RE, Trung HD, Roelants P, Backeljau T, Coosemans M (2001) Confirmation of *Anopheles varuna* in vietnam, previously misidentified and mistar‐ geted as the malaria vector *Anopheles minimus*. Am. J. Trop. Med. Hyg. 65: 729-732.

[3] World Health Organization (2007) Anopheline species complexes in South and South-East Asia. SEARO Technical Publication No. 57. WHO Regional Office for

[4] Singh OP, Nanda N, Dev V, Bali P, Sohail M, Mehrunnisa A, Adak T, Dash AP (2010) Molecular evidence of misidentification of *Anopheles minimus* as *Anopheles fluviatilis*

[5] Harbach RE (2012) Genus *Anopheles* Meigen, 1818. Available: http://mosquito-taxo‐ nomic-inventory.info/genus-anopheles-meigen-1818. Accessed 2012 May 14.

[6] Kanda T, Takai K, Chiang GL, Cheong WH, Sucharit S (1981) Hybridization and some biological facts of seven strains of the *Anopheles leucosphyrus* group (Reid, 1968).

[7] Baimai V, Andre RG, Harrison BA, Kijchalao U, Panthusiri L (1987) Crossing and chromosomal evidence for two additional sibling species within the taxon *Anopheles dirus* Peyton and Harrison (Diptera: Culicidae) in Thailand. Proc. Entomol. Soc.

[8] Baimai V, Poopittayasataporn A, Kijchalao U (1988) Cytological differences and chro‐ mosomal rearrangements in four members of the *Anopheles dirus* complex (Diptera:

[9] Sawadipanich Y, Baimai V, Harrison BA (1990) *Anopheles dirus* species E: Chromoso‐ mal and crossing evidence for another member of the Dirus complex. J. Am. Mosq.

[10] Subbarao SK (1998) Anopheline species complexes in South-East Asia. WHO. Tech.

[11] Peterson HE (1980) A comment on "mate recognition systems". Evolution 34:

[12] Green CA, Rattanarithikul R, Charoensub A (1992) Population genetic confirmation of species status of the malaria vectors *Anopheles willmori* and *An. pseudowillmori* in Thailand and chromosome phylogeny of the Maculatus group of mosquitoes. Med.


[39] Da Silva AN, Dos Santos CC, Lacerda RN, Santa Rosa EP, De Souza RT, Galiza D, Sucupira I, Conn JE, Póvoa MM (2006) Laboratory colonization of *Anopheles aquasalis* (Diptera: Culicidae) in Belém, Pará, Brazil. J. Med. Entomol. 43: 107-119.

[26] Kanda T (1979) Improved techniques for the preparation of polytene chromosomes

[27] Mourya DT, Ilkal MA, Mishra AC, Jacob PG, Pant U, Ramanujam S, Mavale MS, Bhat HR, Dhanda V (1989) Isolation of Japanese encephalitis virus from mosquitoes col‐ lected in Karnataka state, India during 1985 to 1987.Trans. R. Soc. Trop. Med. Hyg.

[28] George S, George JP, Rao JA (1987) Isolation of Japanese encephalitis and West Nile viruses from mosquitoes collected in Kolar district of Karnataka state during 1977-79.

[29] Thenmozhi V, Rajendran R, Ayanar K, Manavalan R, Tyagi BK (2006) Long-term study of Japanese encephalitis virus infection in *Anopheles subpictus* in Cuddalore dis‐

[30] Chakravarty SK, Sarkar JK, Chakravarty MS, Mukherjee MK, Mukherjee KK, Das BC, Hati AK (1975) The first epidemic of Japanese encephalitis studied in India-virologi‐

[31] Choochote W, Sucharit S, Abeyewickreme W (1983) A note on adaptation of *Anophe‐ les annularis* Van Der Wulp, Kanchanaburi, Thailand to free mating in a 30 x 30 x 30

[32] Choochote W, Sucharit S, Khamboonruang C, Somboon P, Maleewong W, Suwapanit P (1985) Adaptation of various species and strains of *Anopheles* mosquitoes to natural

[33] Baker RH, French WL, Kitzmiller JB (1962) Induced copulation in *Anopheles* mosqui‐

[34] Ow Yang CK, Sta Maria FL, Wharton RH (1963) Maintenance of a laboratory colony

[35] Chomcharn Y (1979) The study of *Plasmodium falciparum* in Thailand with respect to the mosquito vector, *Anopheles balabacensis* baisas and the effect of primaquine on the

[36] Gerberg EJ, Barnard DR, Ward RA (1994) Procedures for laboratory rearing of specif‐ ic mosquitoes, In: Manual for mosquito rearing and experimental techniques. Louisi‐

[37] Bangs MJ, Soelarto T, Barodji, Wicaksana BP, Boewono DT (2002) Colonization of *Anopheles maculatus* from Central Java, Indonesia. J. Am. Mosq. Control Assoc. 18:

[38] Kim SJ, Choochote W, Jitpakdi A, Junkum A, Park SJ, Min GS (2003) Establishment of a self-mating mosquito colony of *Anopheles sinensis* from Korea. Korean J. Entomol.

gametocytes and sporogony. Ph.D. Thesis, Mahidol University. pp. 46-48.

copulation in a 30 cm cube cage. J. Parasit. Trop. Med. Ass. Thailand 8: 44-47.

trict, Tamil Nadu, South India. Trop. Med. Int. Hlth. 11: 288-293.

cm cage. Southeast Asian J. Trop. Med. Public Health, 14: 559-560.

of *Anopheles maculatus* by artificial mating. Mosq. News 23: 34-35.

for some *Anopheles* mosquitoes. Mosq. News 39: 568-574.

83: 550-552.

Indian J. Med. Res. 85: 235-238.

76 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

toes. Mosq. News 22: 16-17.

ana: Allen Press. pp. 41-45.

359-363.

33: 267-271.

cal studies. Indian J. Med. Res. 63: 77-82.


[66] Park MH, Choochote W, Kim SJ, Somboon P, Saeung A, Tuetan B, Tsuda Y, Takagi M, Joshi D, Ma Y, Min GS (2008) Non-reproductive isolation among four allopatric strains of *Anopheles sinensis* in Asia. J. Am. Mosq. Control Assoc. 24: 489-495.

[55] Smitinand T (1980) Thai plant names: Botanical names-ver-nacular names. Royal For‐

[56] Jitpakdi A, Choochote W, Insun D, Tippawangkosol P, Keha P, Pitasawat B (1999) Screening of ten plant species for metaphase chromosome preparation in adult mos‐ quitos (Diptera: Culicidae) using an inoculation technique. J. Med. Entomol. 36:

[57] Choochote W, Pitasawat B, Jitpakdi A, Rattanachanpichai E, Riyong D, Leemingsa‐ wat S, Wongkamchai S (2001) The application of ethanol-extracted *Gloriosa superba* for metaphase chromosome preparation in mosquitoes. Southeast Asian J.Trop. Med.

[58] Choochote W, Rongsriyam K, Pitasawat B, Jitpakdi A, Rattanachanpichai E, Junkum A, Tuetun B, Chaiwong P (2004) Evaluation of the colchicine-like activity of *Gloriosa superba*-extracted fractions for mosquito (Diptera: Culicidae) cytogenetic study. J.

[59] Baimai V, Rattanarithikul R, Kijchalao U (1993) Metaphase karyotypes of *Anopheles* of Thailand and Southeast Asia: I. The Hyrcanus group. J. Am. Mosq. Control Assoc. 9:

[60] Baimai V, Rattanarithikul R, Kijchalao U (1995) Metaphase karyotypes of *Anopheles* of Thailand and Southeast Asia: IV, The Barbirostris and Umbrosus species groups, subgenus *Anopheles* (Diptera: Culicidae). J. Am. Mosq. Control Assoc. 11: 323-328. [61] White GB, Coluzzi M, Zahar AR (1975) Review of cytogenetic studies on anopheline

[62] Choochote W, Jitpakdi A, Sukontason K, Chaithong U, Wongkamchai S, Pitasawat B, Jariyapan N, Suntaravitun T, Rattanachanpichai E, Sukontason K, Leemingsawat S, Rongsriyam Y (2002) Intraspecific hybridization of two karyotypic forms of *Anopheles vagus* (Diptera: Culicidae) and the related egg surface topography. Southeast Asian J.

[63] Park SJ, Choochote W, Jitpakdi A, Junkum A, Kim SJ, Jariyapan N (2003) Evidence for a conspecific relationship between two morphologically and cytologically differ‐

[64] Choochote W, Jitpakdi A, Rongsriyam Y, Komalamisra N, Pitasawat B, Palakul K (1998) Isoenzyme study and hybridization of two forms of *Anopheles sinensis* (Dip‐ tera: Culicidae) in Northern Thailand. Southeast Asian J. Trop. Med. Public Health,

[65] Min GS, Choochote W, Jitpakdi A, Kim SJ, Kim W, Jung J, Junkum A (2002) Intraspe‐ cific hybridization of *Anopheles sinensis* (Diptera: Culicidae) strains from Thailand

ent forms of Korean *Anopheles pullus* mosquito. Mol. Cells 16: 354-360.

est Department, Bangkok.

78 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Public Health, 32: 76-82.

Med. Entomol. 41: 672-676.

vectors of malaria. WHO/MAL/75, 489. 35 p.

Trop. Med. Public Health, 33 (suppl 3): 29-35.

and Korea. Mol. Cells 14: 198-204.

892-895.

59-67.

29: 841-848.

