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Land Use Change and the Microbial Ecology of Anopheles gambiae✔

✔

Thomas M. Gilbreath III, Orange County Vector Control District13001Garden Grove Blvd, Garden Grove, California 92843, tgilbrea@uci.edu, tel:+1(949)943-9720

Guofa Zhou, Program in Public Health,University of California, Irvine3501 Hewitt Hall, Irvine, California 92697, zhouga@uci.edu, tel:+1(949)824-0249

Eliningaya J. Kweka, Tropical Pesticides Research Institute, P.O.Box 3024, Arusha, Tanzaniapat.kweka@gmail.com, tel:+255 754 368748

Guiyun Yan, Program in Public Health,University of California, Irvine3501 Hewitt Hall, Irvine, California 92697, guiyuny@uci.edu, tel:+1(949)824-0175

✔

Thomas M. Gilbreath III

Land Use Change and the Microbial Ecology of Anopheles gambiae

Thomas M. Gilbreath III1, Guofa Zhou

1, Eliningaya J. Kweka

2, and Guiyun Yan

1

1Program in Public Health, University of California, Irvine, CA 92697, USA

2Tropical Pesticides Research Institute, Arusha, Tanzania

Recent studies have suggested that land use changes, such as deforestation, strongly

enhance the productivity of malaria vectors, and thus malaria transmission. However, the

mechanism for habitat productivity enhancement by deforestation is not clear. The present study

conducted a metagenomics analysis of Anopheles gambiae mosquitoes to determine the impacts

of deforestation on bacterial and algal diversity of the larval habitats and subsequent habitat

productivity. The metagenomic analysis was based on high-throughput next-generation

pyrosequencing of microbial 16S and 23S DNA directly extracted from field-collected water and

mosquito specimens to provide a comprehensive view of microbial community diversity. Results

of habitat productivity in different land use scenarios demonstrated a significant effect of land

cover on larval development and adult mosquito productivity. Metagenomic analysis of algal

communities by pyrosequencing of 23S DNA revealed a selective feeding on green algae species

(Chlorophyta), which were far more abundant in the larval gut when compared to the available

potential resources. Quantitative PCR demonstrated that although shaded habitats had higher

bacterial abundance, larval development was poor in these habitats, whereas in sunlit habitats,

bacterial abundance was lower, but larval survivorship was higher and development time was

faster, suggesting algae is a more important food resource for An. gambiae larvae relative to

bacteria. This study provided strong evidence that algal community shifts resulting from

deforestation enhanced habitat productivity for An. gambiae mosquitoes. The implications of

microbial community dynamics in the scope of malaria vector control are discussed.

1

Introduction

Over the past three decades, as a result of increasing population and agricultural

development, the western Kenya highlands have undergone considerable environmental changes

(Himeidan and Kweka 2012). Environmental changes, such as temperature, humidity and

habitat availability can significantly affect mosquito abundance, which in turn affect malaria

transmission intensity (Minakawa, Munga et al. 2005). Past studies have suggested that land use

changes, such as deforestation, strongly enhance the productivity of malaria vectors, and thus

malaria transmission (Walsh, Molyneux et al. 1993). This is because deforestation exposes

aquatic habitats to sunlight, resulting in increased water temperatures . Further, exposure to

sunlight may induce changes in the microbial communities that mosquito larvae use for nutrition.

However, there is little knowledge of how microbial communities in aquatic habitats are

regulated by exposure to sunlight and organic nutrients, and how mosquito larvae respond to

microbial community changes in larval habitats (Merritt, Craig et al. 1992; Merritt, Dadd et al.

1992).

The larval ecology of Anopheles gambiae sensu stricto (An. gambiae), the primary

malaria vector in sub-Saharan Africa, remains poorly understood with respect to feeding

preferences and behaviors (Merritt, Dadd et al. 1992); however, it is known that variability in

aquatic habitat characteristics contributes to significant differences in larval abundance

(Minakawa, Munga et al. 2005). Several studies have demonstrated that larval development is

density dependent, suggesting that available food resources may also play a role in regulating

larval habitat productivity (Gimnig, Ombok et al. 2002; Kweka, Zhou et al. 2012). Previous

work has demonstrated the importance of algae in the growth and development of larvae

(Coggeshall 1926; Howland 1930; Kaufman, Wanja et al. 2006), and others have also shown a

reduction in algal abundance in the presence of larvae (Gimnig, Ombok et al. 2002).

2

Associations between larvae and green algae species (Chlorophyta; Chlorophyceae) have also

been observed (Tuno, Githeko et al. 2006). The surface microlayers (SuM) of aquatic habitats

typically accumulate high numbers of bacteria, algae, and dissolved organic matter (Merritt,

Dadd et al. 1992). Mosquito larvae of the genus Anopheles are typically oriented horizontally

along the air to water interface, and they do the majority of their feeding in the surface

microlayer of their aquatic habitats (Merritt, Craig et al. 1992).

The current study examined how chemical and physical characteristics of larval habitats

under variable canopy coverage affect larval habitat productivity, and assessed the importance of

canopy coverage independent of temperature. The mechanisms of larval population regulation

have recently received renewed interest (Himeidan and Kweka 2012). We hypothesize that An.

gambiae are largely regulated by physical habitat characteristics and the resultant availability of

nutritional resources. We utilized two experiments to further test the hypothesis that reduced

canopy coverage, with or without variation in temperature, would lead to increased larval habitat

productivity due to the resulting increase of photosynthetic microbes that An. gambiae use for

food. A third experiment was conducted to test the effects of food addition in all experimental

treatments. One important technique we used was the metagenomic analysis of the larval habitats

and An. gambiae mosquito larvae, which offers a comprehensive view of microbial communities

(Wooley, Godzik et al. 2010; Wang, Gilbreath et al. 2011). This approach was based on the high-

throughput next-generation pyrosequencing of microbial 16S and 23S DNA extracted directly

from field-collected water and mosquito specimens, thus avoiding the bias associated with

inability to culture some bacteria or algae required by the traditional methods for detecting and

identifying microbes (Wooley, Godzik et al. 2010). The metagenomics approach provides a more

powerful tool to test the hypothesis that larval gut contents would demonstrate preferential

3

feeding by larvae upon the photosynthetic microbes in the surface microlayer of their aquatic

habitats.

Methods

We established an array of artificial habitats, or microcosms, in Iguhu, Kakamega

District, western Kenyan (elevation 1,500m). Three sub-sites were chosen for deforested, semi-

forested and forested habitats (un-shaded, partially shaded and heavily shaded, respectively).

Microcosms for experiment one were constructed using 40cm diameter washtubs, and

approximately two liters of top-soil from the forest edge were added to each. Eight microcosms

were distributed at each sub-site. An. gambiae eggs were transported to the study site and

hatched onsite. One hundred first instar larvae were added into each microcosm. Habitats were

monitored for survivorship and life table data (instar proportions) daily, and adult wing length

was measured. Temperature was logged hourly throughout the duration of the experiment using

HOBO TidbiT water temperature loggers (Onset Computer Corporation, Bourne, MA, USA).

Pooled 50 ml surface water samples (< 2 mm) were taken weekly from each sub-site in triplicate

using a needle and syringe, centrifuged at 10,000 g for 20 minutes, and the resultant pellet was

used for DNA extraction. For each canopy coverage level, triplicate samples of eight larval guts

were removed under a dissecting microscope using sterilized dissection devices.

Temperature-controlled microcosms were constructed using perforated baskets lined with

thin (1 mil) polyethylene plastic sheeting to allow for maximum heat exchange between

microcosms and pools. Replicates were assigned shade or sunlit treatment to mimic forested and

deforested habitats. Shades were constructed with wooden frames and 1.5 mil opaque black

plastic, All microcosms were placed into approximately 4 x 2 x 0.5 meter pools that were filled

with stream water to match the water level in the microcosms.

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. DNA was extracted from larval guts and water sample pellets using the UltraClean Soil

DNA Extraction Kit (Mo Bio Laboratories, Carlsbad, CA, USA). To characterize the bacterial

communities, the variable 16S rDNA region V1-3 was amplified. Algal communities were

characterized using the Domain V of the 23S plastid rRNA gene (Sherwood, Chan et al. 2008).

16S PCR, 23S PCR and pyrosequencing were conducted at Research and Testing Laboratory

(Lubbock, TX) using the Roche Titanium 454 FLX pyrosequencing platform as described

previously (Callaway, Dowd et al. 2010). Sequences were deposited in the NCBI sequence reads

archive (accession number SRA051793). The Ribosomal Database Project (RDP) classifier was

used to assign taxonomic rank with a confidence threshold of 80%

(http://rdp.cme.msu.edu/classifier/classifier.jsp) (Wang, Garrity et al. 2007). We used BLAST

searches to match algal sequences from water and gut samples with known organisms in the

National Center for Biotechnology Information database.

Relative ratios of bacteria were determined with qPCR using bacteria universal

quantitative primers on triplicate samples from the water surface microlayer of habitats. Serial

dilutions of known concentrations of Pseudomonas aeruginosa DNA were used to generate

reference samples in order to determine relative abundance of bacteria as done previously

(Dowd, Hanson et al. 2010).

Food supplementation with approximately 100mg of Tetramin Fish Food (Spectrum

Brands, Inc., Madison, WI, USA) was added daily to eight additional replicates of all treatments

in experiment one and two.

For all experiments, differences in An. gambiae larval survivorship, larval development

time and adult wing length among habitats were compared using analysis of variance (ANOVA)

(JMP Statistical Discovery Software, version 5.1; SAS Institute, Cary, NC). Stepwise

5

regression was used to analyze the effects of environmental variables on larval survivorship and

development time, and indirect effects were analyzed using path analysis and performed by using

SPSS Amos (Amos Development Corporation, Meadville, PA 16335, USA). For pyrosequencing

data, bacterial species richness and diversity analyses were conducted using the program

MOTHUR (Schloss, Westcott et al. 2009). We used MOTHUR to generate matrices of genetic

distance and group sequences into operational taxonomic units (OTUs) For each treatment we

used MOTHUR to calculate Chao1 estimates of richness and ACE estimates of diversity

(Wooley, Godzik et al. 2010).

Results/Discussion

The current study examined the relationship between habitat vector productivity,

microbial food resources and deforestation. Bacterial and algal communities are dependent on

temperature in water and soil, but drastic changes in microbial communities, particularly

photosynthetic microbes, also result from sunlight proliferation to the habitat surface as a

consequence of disforestation. The semi-natural, temperature and light variable treatments

confirm previous findings that larval success is limited by canopy coverage (Afrane, Zhou et al.

2007). Temperature-controlled experiments were designed to control water temperatures while

subjecting habitats to variable sunlight levels. Despite temperatures conducive to high larval

survivorship in the shaded, temperature-controlled habitats, pupation rates were low, presumably

due to the lack of available food resources in the sunlight poor environment (Figure 2). These

conclusions are further supported by the results of food supplementation. When shaded habitats

were supplemented with food, larval performance was nearly identical when compared to food

supplemented sunlit habitats (Figure 2). This study did not examine oviposition preference of An.

gambiae mosquitoes for open sunlit habitats, but it provided strong evidence for enhanced larval

6

development and survivorship as a direct result of microbial community changes in sunlit

habitats.

Of these microbial communities, bacterial and algal communities are thought to be of

most importance (Merritt, Dadd et al. 1992; Kaufman, Wanja et al. 2006). While foraging on

biofilms has been shown to be of particular importance to Aedes albopictus and Ae. aegypti ,

algal abundance has been suggested as an important factor for anophelines (Kaufman, Wanja et

al. 2006; Garros, Ngugi et al. 2008). The current study examined the relative abundance of algae

and bacteria in relation to larval success. Relative bacterial abundance indicated that bacterial

abundance increased with increasing canopy coverage. Even though shaded habitats in

temperature-controlled experiments had higher bacterial abundance, larval development was

poor in these habitats. Further, in sunlit habitats, where lower bacterial abundance was

observed, larval survivorship was higher and development time was faster. Given that these

results were from temperature-controlled microcosms, our study strongly suggest that algae, not

bacteria, is the most important food source for An. gambiae larvae, and habitat productivity is

mostly limited by algal concentration of larval habitats.

The Proteobacteria were detected in highest abundance in all samples from both the water

SuM and LGC; however, a notable shift in Proteobacteria class representation was observed. The

Enterobacteriaceae (Gammaproteobacteria) were shown to be effective colonizers of the larval

gut, and previous work has demonstrated that the adult gut selectively favors the colonization of

Enterobacteriaceae (Wang, Gilbreath et al. 2011) (Figure 1).

The mechanisms for differential ingestion of certain microbial groups by An. gambiae

larvae are not clear. One potential mechanism is larval behavioral modification in response to

resource availability. For example, it has been shown that An. quadrimaculatus larvae employ a

7

suite of behaviors to discriminate between available food resources on or near the surface and in

the water column (Merritt, Craig et al. 1992). Feeding intensity can also be up-regulated by the

presence of phagostimulants produced by microbial sources (Dadd 1970).

Productive anopheline habitats have long been strongly correlated with sunlight and the

presence of algae (Coggeshall 1926; Howland 1930; Gimnig, Ombok et al. 2002; Tuno, Githeko

et al. 2006). Further, Tuno et al. found an association between the green alga, Rhopalosolen

species (Chlorophyta) and An. gambiae abundance and body size (Tuno, Githeko et al. 2006).

Although no Chlorophyta sequences were obtained from the forest water samples, the gut

contents of larvae reared in the forest were dominated by chlorophytes (≈ 84%) (Figure 1). This

distinction is likely due to the very low abundance of chlorophytes in the habitats and further

reduction by larvae. Alternatively, larvae may be employing atypical foraging techniques (i.e.,

scraping, shredding) in the forested habitats with low algal growth. Surprisingly, diatoms rather

than chlorophytes dominated the LGC from the deforested area (Figure 1). This is likely due to

the abundance of photosynthetic food resources in the deforested site and a buildup of diatoms

that are more resistant to digestion in the mosquito gut due to their characteristic cell wall

composed of recalcitrant silica .

Approximately 80% of bacterial sequences were identified to species based on sequence

similarity analysis; however, algae were only identified to the family level (48%) due to the

limited number of sequences available in GenBank. We did not determine the contribution of

individual microbial species to nutrition of mosquito larvae, but the mosquito developmental

data from the temperature-controlled experiments is consistent with the general findings of the

pyrosequencing data: habitats lacking algal growth largely do not support larvae to adulthood.

8

These results demonstrate that sunlight proliferation to habitats and the resultant growth

of photosynthetic microbes are critical to the success of Anopheles gambiae mosquitoes. Further,

this work suggests that larval habitat productivity is up-regulated by the eutrophication of aquatic

habitats associated with agriculture. The combined effect of sunlight proliferation due to canopy

removal and the use of fertilizers contribute to habitat eutrophication and algal blooms that will

be beneficial to larval populations. This study enhances our ability to identify or predict aquatic

habitats suitable for malaria vector production and thus facilitates mosquito larval control

targeted at the most productive habitats.

ACKNOWLEDGEMENTS

We thank Anne Vardo-Zalik and Ming-Cheih Lee for helpful discussion and expertise, and the

two anonymous reviewers for constructive comments. This work was supported by grants from

the National Institutes of Health (R01 AI050243 and D43 TW001505)

REFERENECES

Afrane, Y., G. Zhou, et al. (2007). "Life-table analysis of Anopheles arabiensis in western Kenya

highlands: effects of land covers on larval and adult survivorship." American Journal of

Tropical Medicine and Hygiene 77(4): 660.

Callaway, T., S. Dowd, et al. (2010). "Evaluation of bacterial diversity in the rumen and feces of

cattle fed different levels of dried distillers grains plus solubles using bacterial tag-

encoded FLX amplicon pyrosequencing." Journal of Animal Science 88(12): 3977.

Coggeshall, L. T. (1926). "Relationship of plankton to anopheline larvae." American Journal of

Epidemiology 6(4): 556-569.

9

Dadd, R. (1970). "Relationship between filtering activity and ingestion of solids by larvae of the

mosquito Culex pipiens: A method for assessing phagostimulant factors." Journal of

Medical Entomology 7(6): 708-712.

Dowd, S. E., J. D. Hanson, et al. (2010). "Survey of Fungi and Yeast in Polymicrobial Infections

in Chronic Wounds." Journal of Wound Care 20(1): 40-47.

Garros, C., N. Ngugi, et al. (2008). "Gut content identification of larvae of the Anopheles

gambiae complex in western Kenya using a barcoding approach." Mol. Ecol. Resour.

8(3): 512-518.

Gimnig, J., M. Ombok, et al. (2002). "Density-dependent development of Anopheles gambiae

(Diptera: Culicidae) larvae in artificial habitats." Journal of medical entomology 39(1):

162-172.

Himeidan, Y. E. S. and E. Kweka (2012). "Malaria in east African highlands during the past 30

years: impact of environmental changes." Frontiers in Systems Biology 3: 315.

Howland, L. J. (1930). "Bionomical investigation of English mosquito larvae with special

reference to their algal food." The Journal of Ecology: 81-125.

Kaufman, M., E. Wanja, et al. (2006). "Importance of algal biomass to growth and development

of Anopheles gambiae larvae." Journal of medical entomology 43(4): 669-676.

Kweka, E. J., G. Zhou, et al. (2012). "Effects of co-habitation between Anopheles gambiae ss

and Culex quinquefasciatus aquatic stages on life history traits." Parasit. Vectors 5(1): 33.

Merritt, R., D. Craig, et al. (1992). "Interfacial feeding behavior and particle flow patterns

ofAnopheles quadrimaculatus larvae (Diptera: Culicidae)." Journal of Insect Behavior

5(6): 741-761.

10

Merritt, R. W., R. H. Dadd, et al. (1992). "Feeding behavior, natural food, and nutritional

relationships of larval mosquitoes." Annual Review of Entomology 37(1): 349-374.

Minakawa, N., S. Munga, et al. (2005). "Spatial distribution of anopheline larval habitats in

Western Kenyan highlands: effects of land cover types and topography." American

Journal of Tropical Medicine and Hygiene 73(1): 157-165.

Schloss, P., S. Westcott, et al. (2009). "Introducing mothur: open-source, platform-independent,

community-supported software for describing and comparing microbial communities."

Applied and environmental microbiology 75(23): 7537.

Sherwood, A. R., Y. L. Chan, et al. (2008). "Application of universally amplifying plastid

primers to environmental sampling of a stream periphyton community." Molecular

Ecology Resources 8(5): 1011-1014.

Tuno, N., A. Githeko, et al. (2006). "The association between the phytoplankton, Rhopalosolen

species (Chlorophyta; Chlorophyceae), and Anopheles gambiae sensu lato (Diptera:

Culicidae) larval abundance in western Kenya." Ecological Research 21(3): 476-482.

Walsh, J., D. Molyneux, et al. (1993). "Deforestation: effects on vector-borne disease."

Parasitology 106(Suppl 1): S55-75.

Wang, Q., G. M. Garrity, et al. (2007). "Naive Bayesian classifier for rapid assignment of rRNA

sequences into the new bacterial taxonomy." Applied and Environmental Microbiology

73(16): 52-61.

Wang, Y., T. M. Gilbreath, et al. (2011). "Dynamic Gut Microbiome across Life History of the

Malaria Mosquito Anopheles gambiae in Kenya." PloS one 6(9): e24767.

Wooley, J. C., A. Godzik, et al. (2010). "A primer on metagenomics." PLoS Computational

Biology 6(2): e1000667.

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FIGURE 1. Bacterial (A) and algal (B) diversity at the phylum level revealed by

pyrosequencing. Major bacterial phyla are presented and remaining taxa and unclassified

bacteria are pooled and referred to as Other. SuM: water surface microlayer; LGC: third-instar

larval gut contents.

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FIGURE 2. Chlorophyll a concentration (A) and pupation rates (B) of Anopheles gambiae in

temperature controlled microcosms, with and without food supplementation. Food

supplementation in shaded microcosms resulted in nearly identical pupation rates to sunlit

microcosms (P = 0.93). Food supplementation also resulted in increased chlorophyll a and larval

pupation rates in all semi-natural treatments.

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