• Anopheles albimanus;
  • An. vestitipennis;
  • An. darlingi;
  • fatty acids (FA);
  • larval habitats


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Larvae of the three important Central American malaria vectors, Anopheles albimanus, An. vestitipennis, and An. darlingi, are found in distinctly different habitats broadly defined by hydrology and aquatic vegetation, but little is known about the actual food quality and quantity of these habitats. Polyunsaturated fatty acids (PUFA) are of special interest, because mosquitoes require 20:5ω3 (EPA), 20:4ω6 (ARA), and 22:6ω3 (DHA) and without an adequate supply of these PUFAs they are not able to complete their life cycle. We collected samples of larvae and their corresponding habitats and analyzed their fatty acid (FA) composition to reveal if there are any species-specific and habitat-specific differences in FA composition, and if habitat FA differences can be linked to differences in the mosquito FA pattern and, ultimately, mosquito performance. We also assessed how FA of wild larvae compare to the laboratory-reared larvae. Habitats were generally low in essential PUFAs and there were no significant differences among the FA composition of habitat samples. There were significant differences in FA composition of larvae. An. darlingi contained significantly higher amounts of FA, specifically a higher content of ω-6 PUFA, represented mainly by the linoleic acid (18:2ω-6). Large differences were found between field-collected and laboratory-reared An. vestitipennis larvae, especially in the content of PUFAs. The laboratory-reared larvae contained significantly more of the total FA, ω3 PUFA, and MUFA. The laboratory-reared larvae contained three to five times more essential PUFAs, EPA, and DHA. However, there were no differences in the total dry weight of the 4th instar larvae between the wild vs laboratory-reared larvae. Total FA in both larvae and habitats of An. albimanus and An. darlingi were positively correlated with the concentration of particulate organic carbon and nitrogen (POC, PON) in their respective habitats, but no such correlation was found for An. vestitipennis. PUFA are a good indicator of nutritional quality, although factors controlling the success of anopheline development from larval habitats are likely to be more complex and would include the presence of predators, pathogens, and toxins as interacting factors.


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Anopheline mosquitoes, specifically Anopheles darlingi (Root), An. albimanus (Wiedemann), and An. vestitipennis (Dyar and Knab) are of special interest as malaria vectors in Central America. Larvae of these species are generally found in distinctly different habitats broadly defined by hydrology and aquatic vegetation (Manguin et al. 1996, Rejmánková et al. 1993, 1998). Although Anopheles habitats were recognized and their environmental determinants defined, so far no attempt has been made to link these environmental determinants to the food availability for larvae. Except for a study on bacteria counts in these different types of habitat (Rejmánková et al. 2000), little is known about the actual food quality and quantity. It has been shown through reciprocal transplant experiments that each species performs best in its own habitat (Grieco et al. 2007).

Anopheline larvae use their lateral palatal brushes to collect bacteria and organic particles of 1.5 to 4.5μm in diameter from the water surface microlayer (Walker and Merritt 1993). Compared to subsurface water, the surface microlayer contains relatively high amounts of nutrients, organic material both particulate and dissolved, and various microorganisms (bacteria, protozoans, amoebae). These live and dead organic materials provide the necessary dietary items, proteins, lipids, and carbohydrates for larval nutrition. Lipids are an important food component for mosquito larvae because they provide a concentrated and in-osmotic form of energy storage and a source of essential biochemical nutrients.

Fatty acid (FA) constituents of lipids are present in a great structural variety, and are thus increasingly being used as chemical markers of biogeochemical processes and trophic relationships (Desvilettes et al. 1997, Napolitano 1999, Müller-Navarra et al. 2004, Marazouzi et al. 2008, Arts et al. 2009). While the saturated palmitic acid (16:0) is often one of the most abundant fatty acids in lipid extracts, the interest of nutritional studies has concentrated on polyunsaturated fatty acids (PUFA) with two or more double bonds (Parrish 1999). Food quality differences can be related to the content of certain PUFAs (Müller-Navarra 1995, Ahlgren et al. 1997, Müller-Navarra et al. 2000). Some of these PUFAs are essential to the normal functioning of the cells and they or their corresponding precursors have to be obtained in animal diets. In most animals, the 18-carbon chain, 18C, PUFAs can be converted to the longer-chain essential PUFAs, specifically 18:2(ω–6) to 20:4(ω–6) – arachidonic acid, ARA, and 18:3(ω–3) to 20:5(ω–3), eicosapentaenoic acid, EPA, and further to 22:6(ω–3) – docosahexaenoic acid, DHA. Mosquitoes seem to be an exception because their dietary FA requirements cannot be satisfied by the C-18 PUFAs (Dadd 1981, 1983, Dadd et al. 1987). They require some 20- and 22-C polyunsaturated fatty acids (PUFA), specifically 20:5ω3 (EPA), 20:4ω6 (ARA), and 22:6ω3 (DHA) and without an adequate supply of these PUFAs they are not able to fly (Dadd et al. 1987, 1988). The presence of proper PUFAs is essential to all artificial diets, especially when mass-rearing of mosquito is considered (Damiens et al. in press). Adult females may get these from a blood meal (Nor Aliza and Stanley 1998) but these PUFAs are believed essential in the larval stage for flight muscle development. A detailed list of FA from container habitats of Aedes triseriatus is provided in Kaufman et al. (1999) but similar information for anopheline habitats has not been available.

To reveal the importance of feeding habitats for the nutrition of anopheline larvae, we analyzed the FA composition of larvae of three malaria transmitting mosquito species and their corresponding habitats. We were looking for a specific marker or FA pattern that would indicate habitat suitability for a particular species. This included the following questions: Are there any species-specific differences in larval FA composition among anopheline larvae? Are there any seasonal differences in larval composition? How do FA profiles of wild larvae compare to the laboratory-reared larvae? What are the differences in habitat composition? Can habitat FA differences be linked to differences in the mosquito FA pattern and, ultimately, mosquito performance? We also aimed to solve some methodological issues, specifically how to best collect samples for the habitat analysis.

An understanding of the spatial and temporal distribution of dietary resources available to mosquito larvae is needed in order to clarify the relationship among food availability, vector competence, and mosquito fitness. Not only does the nutrient availability within the habitat have to meet a minimum dietary requirement for proper larval development, but the food consumed in the larval stage is critical for a number of physiological processes that impact adult performance (Timmermann and Briegel 1996). It is conceivable that the dependence on blood meals developed in response to insufficient essential biochemical nutrients, e.g., certain PUFAs in the larval habitat. Our contribution provides the first information on PUFAs in anopheline larvae and their natural habitats and can serve as a basis for future trophic/nutritional studies.


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Larval and habitat sample collections

This paper focuses on three malaria vectors, An. albimanus, An. vestitipennis, An. darlingi, and a non-vector species, An. crucians. We collected samples from various locations throughout northern and central Belize covering the dry season (January – May) and wet season (July – November). Anopheles albimanus is the most frequently occurring (albeit the least efficient) malaria vector in Belize. The larvae and habitat samples were collected in its most typical and frequent habitat – shallow marshes with sparse emergent macrophytes (Eleocharis spp.) and floating mats of cyanobacteria. Anopheles crucians frequently occurs in similar marsh types but with somewhat denser macrophytes. Anopheles vestitipennis habitats are limited to marshes with tall dense macrophytes (Typha domingensis, Cladium jamaicense) and flooded forests. Detritus assemblages in rivers, particularly tributaries of the Sibun River, contained habitats of An. darlingi.

Larvae, mostly 3rd and 4th instars, were collected with a standard dipper, identified (live) in the laboratory, rinsed in distilled water, and stored in liquid nitrogen. During both sampling periods we collected An. vestitipennis, An. albimanus, and An. crucians (Wiedemann). Larval stages of An. darlingi are sometimes rather difficult to find and we were not able to collect enough samples of this species during the dry season. In the wet season we found enough suitable habitats of An. darlingi to include this important malaria vector in the analysis. Since An. crucians is not a malaria vector, we included information on its larval and habitat FA composition in the appendices but did not include it in the comparative data analysis.

The samples from larval habitats (“habitat samples” from now on) were collected in two ways: (1) a dipper was lowered just below the water surface (∼ 0.5 cm) and moved very slowly to collect about 250 ml of surface water. Known volumes of each water sample, usually 120 ml, were then filtered through a pre-combusted glass-fiber filter (Whatman-GF/C, 2.5 cm diameter), the remaining sample was processed for analyzing particulate organic carbon and nitrogen (see below); (2) large pre-combusted GF/C filters (15 cm diameter) were gently placed on the water surface and then lifted. The water collected with the large filters was assumed to correspond to the food available in the surface microlayer. The volume of water collected with large filters was 8 ml, which corresponded to an approximate 450 μm thick surface layer. All samples were kept in liquid nitrogen until they were transported to the laboratory to be stored in a deep freezer (–80° C) until the analysis.

Experimental procedure for laboratory-reared Anopheles vestitipennis

Female An. vestitipennis were collected in human-baited collections at a field station located in Orange Walk Town, Belize. Eggs from blood-engorged populations of An. vestitipennis were set in enamel pans filled to a depth of 4.4 cm with rain water. To prevent crowding, no more than 200 larvae were kept in a single pan. Larvae were fed a standard diet of ground fish flakes (Tetramin#63194;) and brewer's yeast mixed in a 3:1 (weight/weight) ratio. The temperature in the insectary ranged from 29–32° C, with a relative humidity of 87–90%, and a photoperiod of 13:11 L:D. For FA analyses, we collected three replicate samples for each life stage, except for the 1st instar where a small size did not allow for replicates.

Fatty acid analysis

A slightly modified method of Kattner and Fricke (1986) was used for fatty acid analyses. After freeze-drying, samples were weighed (Mettler H31AR analytical balance), crushed in combination with an internal standard (heneicosanoic acid, 21:0), and extracted with dichlormethane:methanol (2:1) in the refrigerator. The first extraction took place overnight. The following morning, the extract was removed to new test tubes and the sample was extracted in the original test tube in dichlormethane:methanol (2:1) for two to three h, followed by the third extraction that lasted half an hour to one h. After the last extraction step, the extraction solvent was evaporated off using an N-evaporation unit. For methylation, a 3% solution of sulfuric acid in methanol and n-hexane was added. The test tubes were tightly closed, placed on a heating block, and heated at 80° C for at least 4 h. After methylation, they were cooled and 8 ml of HPLC grade water was added and the liquid was transferred to pear-shaped flasks. Test tubes were rinsed with 5 ml of n-hexane and a rinse was added to the flasks. After shaking, the flasks were set so that the two phases could separate. The hydrophilic phase on the bottom of the flasks was removed by Pasteur pipette and transferred to new flasks and fresh n-hexane was added. This phase-separation was repeated three times to quantitatively extract the fatty acid methyl ester. N-hexane from the lipophilic phase was then evaporated under nitrogen. Samples were transferred to amber vials with Pasteur pipettes after the addition of 1 ml of n-hexane and again evaporated under nitrogen and dissolved in an exact amount of n-hexane. Vials containing samples were stored at –80° C until they could be analyzed. Methyl esters extracted from the FA were analyzed using gas chromatograph (Hewlett Packard 6892) equipped with a programmable temperature vaporization – injector (PTV), fused silica DB-Wax capillary column, and a flame ionization detector (FID). Fatty acid methyl-ester peaks were compared with retention times of standard mixes (37 component FAME Mix from Supelco). Peaks were quantified by comparing peak areas with the internal standard, and corrected for response factors determined from quantitative mixes (Alltech). Unidentified peaks were either FA for which no standards can be purchased or lipophilic components other than fatty acids. The fatty acid nomenclature used here is of the form 18:2w6 nomenclature, where “18” designates the total number of carbon atoms, “2” the number of double bonds, and “w6” the position of the first double bond from the methyl end (w) of the molecule. Fatty acid abbreviations used in this article can be found in Table 1.

Table 1.  Fatty acid abbreviations used in this article.
AbbreviationFull nameStructure
FA(s)Fatty acid(s)
SAFA(s)Saturated fatty acid(s)
MUFA(s)Monounsaturated fatty acid(s)
PUFA(s)Polyunsaturated fatty acid(s)
18C PUFA(s)PUFAs with 18 carbon atoms
20–22C PUFA(s)PUFAs with 20–22 carbon atoms
Linoleic acid18:2ω6
ARAArachidonic acid20:4ω6
Docosapentaenoic acid22:5ω6
Linolenic acid18:3ω3
Octadecatetraenoic acid18:4ω3
EPAEicosapentaenoic acid20:5ω3
DHADocosahexaenoic acid22:6ω3
Sumω6Total ω6 FA
Sumω3Total ω3 FA

Samples for particulate carbon and nitrogen

Surface water samples were first screened through a mesh (0.5 mm) to eliminate larger particles and were then filtered directly onto pre-combusted (2 h at 450°C) glass-fiber filters (Whatman GF/C). Carbon and nitrogen content were determined with a CHN analyzer (Europa ANCA elemental analyzer).

Data analysis

ANOVA was used to test the differences in individual FA among the species. To meet the ANOVA model assumption, we log-transformed some of the variables to normalize the data. The effects of POC and PON on the fatty acid content were evaluated using a linear regression carried out in StatView 4.51.


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The habitat FA profiles obtained from both small and large filters (Tables 2 and 3) were dominated by SAFAs, contributing 43 to 54% of all the FA and these consisted mostly of palmitic (16:0) and steraic (18:0) acids. MUFAs accounted for 23 to 30% of all FA with oleic (18:1ω9), 16:1, and 22:1ω9 acids being the most abundant. Linoleic acid (18:2ω6) was the most abundant of the ω6 PUFAs on both small and large filters, followed by 22:2ω6, 20:2ω6, and 20:4:ω6 (ARA) on small filters and 20:2ω6, and 20:4:ω6 (ARA) on large filters. The ω3 series was dominated by linolenic (18:2ω3), 18:4ω3, and 20:3ω3 on both types of filters. There were no significant differences among habitats of individual species.

Table 2.  Proportions of fatty acids as percentages of total identified fatty acids in larval habitats (small filters); wet season, 2001.
Fatty acids An. albimanus (n=16) An. vestitipennis (n=8) An. darlingi (n=7)
ΣSAFA 48.55   52.37   45.11  
18:1ω9 cis8.773.439.764.989.653.18
18:1 ω9 trans3.922.916.827.342.941.26
ΣMUFA 25.55   32.06   24.39  
18:2ω6 trans0.682.160.441.251.982.41
20:4ω6 (ARA)1.811.350.750.661.190.75
20:5ω3 (EPA)2.671.560.790.691.030.89
22:6ω3 (DHA)1.171.360.180.240.450.38
ΣPUFA 25.88   15.56   30.52  
Table 3.  Proportions of fatty acids as percentages of total identified fatty acids from larval habitats (large filters); wet season, 2001.
Fatty acids An. albimanus (n=15) An. vestitipennis (n=3) An. darlingi (n=8)
ΣSAFA 50.92   43.47   54.29  
18:1ω9 cis18.913.8817.810.0811.725.73
18:1 ω9 trans2.811.082.70.834.314.69
ΣMUFA 29.72   28.12   22.76  
18:2ω6 cis3.572.449.536.096.192.45
18:2ω6 trans0.761.89001.822.22
20:4ω6 (ARA)0.860.631.421.110.47
20:5ω3 (EPA)0.50.420.420.181.090.77
22:6ω3 (DHA)0.250.350.
ΣPUFA 19.35   28.39   22.96  

Relative distribution of individual FA (in % of sum FA) between small and large filters was compared for each pair by a linear regression. There was a close correlation among the proportions of individual acids (P<0.0001 in 34 of 38 pairs, P<0.001 in the remaining; r² between 0.5 and 0.9; data not shown). Many large filters were enriched in 18:1ω9 compared to small filters, while many small filters were enriched in 22:1ω9 and 22:2ω6 (Tables 2 and 3).


Total FA in larvae of 3rd and 4th instars was very similar for An. albimanus and An. vestitipennis, 5.34 %±2.43 SD and 5.08 %± 2.01 SD of dry weight, respectively, and was significantly higher for An. darlingi (7.8 %± 1.99 SD of dry weight). In all species, the major contributor among SAFA was palmitic acid (16:0), followed by steraic (18:0) acid. MUFA were mainly represented by oleic acid (18:1ω9) (Table 4). Palmitic and oleic acids each contributed roughly 17–28% of total FA in each species. PUFAs were mainly represented by linoleic acid (18:2ω6) contributing around 7.4 and 12.3 % for An. albimanus and An. vestitipennis, respectively, while it was more abundant in An. darlingi (17.9 %). Palmitic, oleic, and linoleic FAs constituted over 50% of the fatty acid content in all three species. The fourth and fifth most abundant FAs were 18:0 and 16:1, respectively, contributing around 7% each to the total FAs. The variability of individual FAs was large, often higher among larvae of the same species from one habitat than among species from different habitats. Yet there were some consistent differences in mean FA values and FA groups among individual species (Figure 1). Anopheles darlingi larvae contained a significantly higher concentration of total FA and ω6 PUFA (and consequently a significantly lower ω3/ω6 ratio) than the other two species. Higher concentration of linoleic acid (18:2ω6) was the main reason for this higher ω6 PUFA content in An. darlingi (Figure 1). The proportions of the essential PUFAs, EPA, and ARA were similar, each representing between 3% and 4.6% of the total FA (Table 4). However, EPA seemed to be somewhat higher in An. albimanus compared to An. vestitipennis and An. darlingi, while no significant differences were found among the species in the ARA content (Figure 1). This resulted in a significantly higher EPA/ARA ratio in An. albimanus (Figure 1). Quantities of DHA were small in all species, 0.15% to 0.27 % of the total FA (Table 3) with no differences among species (data not shown).

Table 4.  Proportions of fatty acids as percentages of total identified fatty acids from mosquito larvae; wet season, 2001.
Fatty acids An. albimanus (n=21) An. vestitipennis (n=6) An. darlingi (n=8)
ΣSAFA 44.5   37.39   39.53  
18:1ω9 cis19.454.4317.295.3120.091.82
18:1 ω9 trans2.811.034.772.482.350.41
ΣMUFA 31.13   36.16   30.76  
18:2ω6 trans0.190.88000.751.08
20:4ω6 (ARA)3.071.322.851.812.460.81
20:5ω3 (EPA)4.191.742.471.832.390.74
22:6ω3 (DHA)
ΣPUFA 24.4   26.43   29.69  

Figure 1. Sum of fatty acids, sum of ω6 PUFA, ratio ω3/ω6 PUFA, ratio EPA/ARA, and individual fatty acids [mg g–1 dry weight] in mosquito larvae that showed significant differences among species; A –An. albimanus, V –An. vestitipennis, D –An. darlingi. Error bars indicate the standard error. Bars sharing the same letter are not significantly different from each other.

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Unidentified peaks (UFA) contributed 8.9%, 12.7%, and 14.1% of total extracted methyl esthers for An. albimanus, An. vestitipennis, and An. darlingi, respectively.

Comparison of FA in individual life stages of laboratory-reared Anopheles vestitipennis

There is no spatial preference within a habitat among individual life stages (instars) of mosquito larvae, i.e., individual life stages of one species do not have a preference for a certain spatial location within a habitat. From our field collections we did not have enough material to compare FA composition of individual instars from the same habitat. However, we were able to use laboratory-reared larvae of An. vestitipennis for this comparison. Table 5 shows the proportions of individual FA from the sum of FA for the four larval instars, pupae, females, and males of laboratory-reared An. vestitipennis. Similar to larvae from the natural habitats, the palmitic acid oleic acid and linoleic acid contributed most to the total FA. The sum of fatty acids in the total biomass increased almost five times from 1st instars to pupae and then dropped by a factor of 1.4–1.6 in adult stages depending on gender (Figure 2). Contrary to the FA, the total dry weight of individual stages increased and was highest in the adults (data not shown). Large differences could be found between field-collected and laboratory-reared An. vestitipennis larvae, especially in the content of PUFAs. The laboratory-reared larvae (combined 3rd and 4th instar) contained significantly more of the sum of FA (73.2 vs 46.8 mg/g; P <0.005), ω3 PUFA (10.1 vs 5.1 mg/g; P <0.005) and ω6 PUFA (13.5 vs 6.9 mg/g; P <0.001). Specifically, the laboratory-reared larvae contained three to five more EPA and DHA than field-collected larvae. The food for laboratory-reared larvae was clearly richer in ω3 PUFA, especially EPA. However, there were no differences in the total dry weight of the 4th instar larvae (laboratory-reared: 0.234 mg per individual; field: 0.229 mg per individual).

Table 5.  Proportion of individual fatty acids as percentages of total fatty acids; laboratory-reared larvae of Anopheles vestitipennis; 1st, 2nd, 3rd, and 4th instars (n = 3; no replicated samples were run for the 1st instar).
Fatty acids1st2nd3rd4thPupaFemaleMale
14:03.04 2.36 0.76 2.14 0.7 2.32 0.33 1.92 0.56 0.69 0.21 1.67 2.06
15:01.23 0.64 0.09 0.58 0.06 0.78 0.28 0.62 0.28 0.31 0.37 0.37 0.51
15:0i1.58 1.19 0.4 1.24 0.28 2.05 1.02 0.92 0.75 1.54 2.41 1.67 2.9
16:020.34 18.93 0.71 20.24 3.13 22.88 2.52 23.58 0.82 26.65 7.14 23.49 7.05
16:0i0.63 0.31 0.13 0.25 0.07 0.28 0.12 0.18 0.19 0.01 0.01 0.08 0.12
17:00.81 0.78 0.2 0.69 0.13 0.83 0.23 0.8 0.26 0.11 0.03 0.28 0.22
17:0i0 0.43 0.13 0.35 0.13 0.5 0.13 0.44 0.3 0.03 0.03 0.07 0.08
17:0a1.03 0.47 0.45 0.27 0.23 0.33 0.31 0.19 0.21 0.18 0.31 0.3 0.5
18:014.65 7.11 1.67 8.23 2.25 6.2 0.76 5.92 0.48 4.42 0.26 4.34 0.62
19:00.93 0.31 0.33 0.16 0.19 0.17 0.1 0.07 0.08 0.18 0.13 0.14 0.12
20:01.05 1.08 0.82 1.09 0.6 0.93 0.37 0.63 0.14 1.06 0.78 1.17 0.66
22:00.33 0.88 0.43 0.78 0.21 0.8 0.12 0.5 0.14 0.65 0.4 0.71 0.21
23:00.49 0.09 0.08 0.27 0.21 0.13 0.03 0.04 0.03 0.01 0.02 0.04 0.04
24:00.39 0.25 0.12 0.22 0.06 0.16 0.03 0.08 0.06 0.11 0.08 0.1 0.07
ΣSAFA 46.5 34.83   36.51   38.36   35.89   35.95   34.43  
14:10 0.08 0.14 0.25 0.22 0.22 0.26 0.17 0.16 0.42 0.35 0.24 0.34
15:10 0 0 0.05 0.06 0 0 0 0 0 0 0.11 0.22
16:14.45 7.6 1.48 6.42 2.73 7.75 1.65 7.52 0.95 14.73 1.4 13.63 3.05
17:10.98 0.77 0.22 0.75 0.13 0.87 0.55 0.78 0.48 0.17 0.04 0.49 0.46
18:1ω9 cis0 16.68 2.58 18.48 1.61 19.06 2.52 18.59 1.21 23.39 1.44 22.28 2.66
18:1 ω9 trans10.74 3.31 0.47 2.58 0.3 2.97 0.69 3.02 0.67 0.87 0.26 1.17 0.95
20:1ω91.79 0.62 0.18 0.56 0.05 0.61 0.27 0.43 0.21 0.55 0.83 0.35 0.38
20:1ω70.52 0.47 0.78 0.11 0.12 0.2 0.13 0.1 0.13 0.21 0.36 0.11 0.19
22:1ω90.26 0 0 0.03 0.02 0.04 0.05 0 0 0.12 0.22 0.12 0.24
24:10 0 0 0.01 0.01 0 0 0 0 0 0 0 0
ΣMUFA 18.74 29.53   29.24   31.72   30.61   40.46   38.5  
18:2ω6cis1.66 14.03 3.94 13.4 2 12.55 2.67 15.07 3.8 8.51 1.51 12.07 0.22
18:2ω6 trans6.87 0.25 0.44 2.02 1.86 0.55 0.59 0.75 0.53 0.97 1.69 0.46 0.31
18:3ω60 0.15 0.13 0.2 0.08 0.39 0.3 0.57 0.43 0.28 0.16 0.2 0.13
18:3ω31.02 2.06 0.11 1.71 0.26 1.49 0.15 1.71 0.31 1.02 0.24 1.48 0.15
18:4ω317.32 5.01 4.4 2.49 0.98 2.13 0.96 1.53 0.51 2.07 2.52 2.64 3
20:2ω61.56 0.83 0.88 1.61 2.32 0.72 0.24 0.8 0.34 1.76 1.77 0.75 0.56
20:3ω60 0.07 0.12 0.38 0.35 0.24 0.22 0.28 0.12 0.28 0.4 0.18 0.27
20:4ω6 (ARA)1.96 2.4 0.67 3.07 2.03 2.47 0.85 2.09 1.41 1.71 0.97 2.19 0.96
20:3ω32.07 0.86 1.18 1.53 1.94 0.76 1.04 0.53 0.67 0.38 0.21 0.6 0.65
20:5ω3 (EPA)1.49 7.94 1.85 6.5 0.63 7.84 1.99 9.44 0.52 6.45 0.85 6.36 3.07
22:2ω60 0 0 0.08 0.13 0.01 0.02 0 0 0 0 0 0
22:6ω3 (DHA)0.82 2.01 0.39 1.25 0.5 0.76 0.1 0.71 0.34 0.13 0.12 0.16 0.13
ΣPUFA 34.77 35.61   34.24   29.91   33.48   23.56   27.09  

Figure 2. The amount of total FA, total ω3 PUFA, and total ω6 PUFA (upper panel) and selected important FA (lower panel) in biomass of individual life stages of laboratory-reared Anopheles vestitipennis (mg per g dry mass). 1- 1st, 2- 2nd, 3–3rd, 4–4th instar, 5- pupa, 6- adult female, 7-adult male. Error bars indicate the standard error.

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Comparison among seasons

For two species, An. albimanus and An. vestitipennis, we had enough material to compare the larval FA content and habitat composition between the dry and wet seasons. Comparisons of habitat samples (small filters) showed An. albimanus habitats to have significantly higher amounts of FA in the dry season, with the total being double of that from the wet season (148.9 and 71 μg/L, respectively). Lower concentration of FA in habitat samples during the wet season most probably reflects the frequent dilution and flushing of habitats by rain. The differences between the seasons for An. vestitipennis habitats were mostly insignificant. This situation is somewhat reflected in larval FA content; the An. albimanus FA amounts were higher in the dry season, while there were no differences for An. vestitipennis larvae.

How does the FA composition of larval tissue reflect habitat composition?

To compare how the larval FA composition corresponds to the FA availability in the habitats we plotted the proportions of individual FA in sum FA from larvae against the proportions in small filters (Figures 3 and 4). For larvae of both An. albimanus and An. vestitipennis, there was a relatively close association between proportions of FA in the larvae and in the corresponding habitats. Figure 4 focuses on PUFAs only. EPA and ARA were always slightly enriched in larval tissue in all species, DHA was always very small, often less than 1%. Of the 18C PUFAs, 18:2w6 was consistently enriched in larval tissue, while 18:4w3 was enriched in habitats. The most significant deviation can be seen in An. darlingi where linoleic acid (18:2ω6) is found in two to three times higher concentrations in larvae as compared to the habitats. The ratio ω3/ω6 was always higher in habitats than in larvae for the corresponding habitat for a given species (habitat – large filters ω3/ω6 ratio = 1.53, 0.72 and 1.07; habitat – small filters = 1.34, 0.94 and 1.27; larvae ω3/ω6 ratio = 0.9, 0.49 and 0.32 for An. albimanus, An. vestitipennis, and An. darlingi, respectively). This indicates the accumulation of ω6 between the two trophic steps. Apparently, linoleic acid (18:2ω6) was mostly responsible for this. Linoleic acid in An. darlingi larvae had 3.32 times higher concentration than in the corresponding habitats, and it was significantly different from a 1:1 ratio (DF= 7; t= 6.455; P= 0.0003). The differences for larvae of the other two species were not as pronounced but still significant: An. albimanus 1.89 times higher (t= 2.429; DF =19; P= 0.025), An. vestitipennis 2.15 times higher (t=4.9; DF = 20; P=0.001).


Figure 3. Proportions of fatty acids in larvae (y-axis) and habitats (x-axis). The line indicates equal proportion. A –Anopheles albimanus, V –An. vestitipennis, D –An. darlingi.

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Figure 4. Proportions of PUFAs in larvae (y-axis) and habitats (x-axis). The line indicates equal proportion; essential PUFAs indicated by full large black dots. A –Anopheles albimanus, V –An. vestitipennis, D –An. darling (note different scales).

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FA vs. POC and PON

In order to further characterize larval habitats we measured the amount of particulate organic carbon (POC) and particulate organic nitrogen (PON) in habitat samples. POC and PON were closely correlated (R2= 0.867; P<0.0001). A box plot shows the comparison of total FA, POC, and PON among the three species as compared to a control (open water) (Figure 5). There were significant positive correlations between both the sum of the FA in larvae and the sum of the FA in the larval habitats, and concentration of both POC and PON in habitats for An. albimanus and An. darlingi, but no such correlation was found for An. vestitipennis (Figure 6).


Figure 5. Box diagram of sums of fatty acids, nitrogen, and carbon concentrations in the habitats of: 1- Anopheles albimanus (A); 2- An. vestitipennis (V); 3- An. darling (D); 4- control. The margins of the boxes enclose the 25th and the 75th percentiles, and the whiskers enclose 10th and 90th percentiles.

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Figure 6. Relationship between particulate organic nitrogen and carbon (mg/L, x-axes) in water and sums of fatty acids in larvae (upper part, mg/g) and habitats (lower part, μg/L). Full circles –An. albimanus, crosses –An. darlingi; squares –An. vestitipennis.

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C/N ratio of 10.5 and 10.8, respectively, did not differ between An. albimanus and An. vestitipennis habitats, but was significantly higher in An. darlingi habitats (15.0). The higher C/N ratio in An. darlingi habitats probably reflects the difference in the quality of prevailing detritus of higher plant origin (= more C) from An. darlingi habitats as compared to mostly algae from habitats of the other two species.


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  7. Acknowledgments


Habitats of the three anopheline species are found in very different environmental settings in terms of hydrology, water quality, and surrounding vegetation (Rejmánková et al. 1993, 1998), although they all were from rather oligotrophic water bodies. An. darlingi is the only species found in rivers, the other species are found either in marshes or in flooded forests, i.e., in stagnant water. The marshes with cyanobacterial mats, An. albimanus habitats, are limited by P and contain little phytoplankton (mean chlorophyll level of 1.75, range 0.05–5.45 μg/ L, unpublished data). While the habitats of An. darlingi and An. vestitipennis are usually well shaded, An. albimanus occurs in sunlit habitats. All habitats have large amounts of detrital material present although the composition for each is quite different. An. darlingi is composed of small twigs, fallen leaves, and seeds from shrubs and trees from the gallery forest along river banks. An. vestitipennis is composed of large quantities of decomposing leaves of marsh macrophytes, mainly Typha domingensis, or leaf litter in the swamp forest. An. albimanus is composed of decaying parts of floating cyanobacterial mats. Because of this large interspecific habitat diversity, we expected to find large differences in the FA composition of individual habitats, but the FA content and composition were surprisingly similar. The proportions of SAFA, MUFA, and PUFA were quite similar to those found in lake seston (Persson and Vrede 2006). Larval habitats had also similar proportions of PUFAs, specifically 18:2ω6 and 18:4ω3, but lower essential PUFAs, EPA, ARA and DHA (Persson and Vrede 2006, Gladyshev et al. 2007). Relatively large proportions of 18:1ω9 and 18:3ω3 indicated a presence of cyanobacteria and green algae, while the fact that we did not find any 16:1ω7 and relatively small proportions of EPA indicated a lack of diatoms (Caramujo et al. 2008, Müller-Navarra 2008).

The amounts of individual FA in both larvae and habitat samples displayed high variation. This can be explained by a very diverse composition of the surface microlayer where the mosquito larvae feed. As a habitat consists of a diverse array of organisms live and/or decomposing, the variability of fatty acids is larger in the habitat than in the larvae, especially in respect to total fatty acids (also see Ahlgren et al. 1997).

The individual FA and the respective totals of habitats of An. albimanus and An. vestitipennis were about 1.5 times higher in the dry season as compared to the rainy season, most likely the result of increased precipitation and subsequent dilution of larval habitat by fresh rain water. This is in contrast to habitats found in temperate climates where the seasonal differences produce dramatic fluctuations in temperature which may result in qualitative differences in FA composition (Marazouzi et al. 2008).

While there was generally a good agreement in the qualitative composition of FA retained by both filters, the quantity of FA was one to two orders of magnitude lower in small filters as compared to the large filters. This reflects the fact that the large filters collect relatively more of the actual surface microlayer, which is enriched in lipids relative to the subsurface water. There was a good agreement between the large and small filters in terms of relative composition of FA. Large filters were enriched in 18:1ω9 compared to the small filters, while many small filters were enriched in 22:1ω9 and 22:2ω6. Enrichment in 18:1ω9 can be interpreted as reflecting a higher proportion of bacteria in the surface microlayer that has been documented previously (Walker and Merritt 1993, Rejmankova et al. 2000, Watton and Preston 2005). The differences in the EPA/ARA ratio between the small and large filters can be also explained by the two sampling methods. There will be relatively more algae in the water column than in the surface layer, and algae are known to have relatively high proportion of EPA. The reason why we did not see the same disproportion in the An. darling habitats can be explained by the fact that these are river habitats, typically shaded and the contribution from algae is less. However when comparing the proportions of these PUFAs between habitats and larvae, there are very similar results for large and small filters.

Because of the similarity of the results, it is hard to recommend which habitat sampling technique is better. The choice becomes more a question of what is easier to manage. Each of the methods used has its associated advantages and disadvantages. While the main advantage of large filters is the collection of samples from the surface microlayer and a higher concentration of a sample, the disadvantage is increased complications due to a more cumbersome handling process of the samples in the field. On the other hand, the small filters are easier to handle, but by using the dipper, one is collecting samples that include both surface and subsurface layers. From a practical perspective, it seems to be easier and more efficient to use the smaller filters.


Anopheline larvae fit well in the range of FA given for aquatic organisms, 2–15% of the dry weight (Ahlgren et al. 1999, Napolitano 1999), and proportions of significant groups of FA in the biomass. Total FA in species analyzed in this paper ranged from 2 to 8% of total dry weight. Also, the relative amounts of individual fatty acids in our larvae lie generally in the range reported for other aquatic invertebrates. In terms of the essential PUFAs, Ahlgren et al. (1999) reported for freshwater invertebrates 8.84%, 3.11%, and 0.19% for EPA, ARA, and DHA, respectively. The ARA and DHA values are comparable to what we found in our larvae, while our EPA values were about half of that given by Ahlgren et al. (1999). The lower values of EPA in both habitats and larvae at our study site may be given by the algal composition. The Belizean habitats are relatively low in diatoms (one of the main sources of EPA; Napolitano 1999) and relatively rich in cyanobacteria (Rejmánková et al. 2004)

Because of their feeding mode, anopheline mosquito larvae filter feed indiscriminately at the water surface, and thus their FA profile should resemble that of their habitats. For a majority of FA it did (Figure 3), the exceptions were the significantly higher amounts of linoleic acid (18:2ω6) in larvae, especially An. darlingi, as compared to their habitats. Larvae apparently concentrate linoleic acid. One of the interesting characteristics of mosquito physiology is their apparent incapability to elongate the 18 C PUFAs, specifically 18:2w6 and 18:3w3 to the PUFAs such as ARA, EPA, and DHA. While most insects can satisfy the essential PUFA requirement with C18 PUFAs, mosquitoes have to obtain them in their food (Dadd and Kleinjan 1979, Dadd 1981, Dadd et al. 1987, Dadd et al. 1988). Without obtaining these essential FAs in their food, the mosquitoes are not capable of completing their development. Even though they may be able to go through metamorphosis to the adult stage, nutritional deficiencies in the larval stage may result in altered physiological processes in the adult such as an inability, or decreased ability, to fly. This is particularly true in anophelines which do not posses the ability to mobilize nutrients from mothers to daughters, and thus the provision of essential PUFAs, as seen in Aedes mosquitoes (Timmermann and Briegel 1996). Experiments conducted by Dadd (1981) resulted in linoleic acid being classified in a group of “semi-active” FA, i.e., those that when fed to larvae were able to support a small proportion of the population to develop adults capable of flying.

Comparison among the wild and reared mosquitoes

Dramatic differences between laboratory-reared organisms and natural populations were reported by Napolitano (1999) and, specifically for mosquitoes, Culex tarsalis (Dadd et al. 1988). Dadd and co-workers examined FA composition in wild-caught adult mosquitoes with those whose larvae were reared on several types of larval food (fish food, rat chow, cat food). The most marked difference was the presence of more than double the amount of linoleic acid (18:2ω6) and less than half the amount of EPA found in the laboratory-reared mosquitoes as compared to wild mosquitoes. Contrary to Dadd et al., we did not find any differences in the linoleic acid content but found much lower EPA values in wild populations as compared to the laboratory-reared larvae. But it is important to point out that when making comparisons one should do so only between similar life stages (Hayashiya and Harwood 1968) and in doing so we found a decrease in the FA content in adult mosquitoes compared to the larvae. This decrease apparently reflects the energy expenditure during metamorphosis (Cavaletto and Gradner 1999).

A number of studies have demonstrated that polyunsaturated fatty acids are required for normal flight in newly emerged mosquitoes (Dadd et al. 1977, Dadd et al. 1988). Mosquito flight, however, is a complex coordination of several physiological processes of which the failure of any one could result in reduced flight activity. An attempt to make a direct association between the fatty acid content of the larval diet and the flight capacity of the resulting adults may be inappropriate. We would, however, suggest future evaluation of the flight indices (Dadd 1983) of newly emerged populations from FA rich and poor habitats as an assessment of the overall fitness of the adult mosquitoes. More detailed evaluations of the adults resulting from FA deficient diets may shed light on how nutrition impacts the fitness of the population and how this may contribute to the overall vector potential of the species.

The concept of compensatory feeding is one that states that an organism can compensate for deficiencies in its diet by altering its feeding behavior (Chapman 1998). This can be accomplished in a number of ways, but researchers have focused on three general paths: 1) switch to feeding on more suitable substrate that maintains a different nutritional balance, 2) adjust the manner in which the nutrients are utilized to maximize efficiency, or 3) adjust the amount of food ingested to compensate for low nutrient levels. Therefore, one possible explanation for how mosquito larvae compensate for low nutrient concentrations in the natural habitat is by increasing their rate of feeding.

The most efficient PUFAs according to Dadd (1981) and Dadd et al. (1988) were EPA, DHA, and ARA. In our samples, DHA was uniformly low, always less than 0.5 % of the total FA. ARA ranged on average from 2.4 to 3.2%, and EPA was in a similar range, from 2.4 to 3.9% (Table 3; small filters). Except for DHA, the other two PUFAs were more concentrated in larvae than in the habitat samples. Dadd's analyses of wild Culex adults was much higher than what we found in a wild population, but their laboratory-reared adults were quite comparable to our laboratory-reared An. vestitipennis. When comparing our results to it has to be kept in mind that Dadd's group analyzed adults, not larval stages. Since the FA content drops in the adults as compared to larvae and pupae (Figure 2), had they presented data for larvae, the differences between their results and ours would be even larger. Dadd's laboratory-reared adults were quite comparable to our laboratory-reared An. vestitipennis. As the laboratory-reared An. vestitipennis displayed much higher PUFA values, it may indicate that our wild populations live in environments relatively poor in PUFA. However, this would be inconsistent with an argument for compensatory feeding. In contrast to Culex, whose larval habitats are generally much more nutrient rich, all of our species were from rather nutrient-poor sites with limited production of green algae.

Anopheles albimanus larvae had a significantly higher ratio of EPA/ARA (Figure 1), which may be a reflection of the habitat (namely small filters) of this species. According to Kuusipalo and Käkelä (2000), the ratio of EPA/ARA is a good indicator of food source (EPA occurs in relatively high concentration in algae, namely diatoms and Cryptophyceae (Desvilettes et al. 1997)), while ARA “accumulates” in higher animals. Two factors may result in a high EPA/ARA ratio: a) higher EPA values, probably reflective of the food source or b) lower ARA values, potentially a combination of a low conversion to ARA and low food values.

Ecological implications

In conclusion, PUFAs are a good indicator of nutritional quality, although factors controlling the success of anopheline development from larval habitats are likely to be more complex and would include the presence of predators, pathogens, and toxins as interacting factors (Kaufman et al. 2006). By better understanding species-specific nutritional requirements, larval foods for laboratory production could be better replicated to ensure more natural adult populations. This becomes vital as more control efforts are focused on the release of genetically modified, laboratory-reared insects. In addition to proving beneficial for countless colonization efforts, a better understanding of mosquito habitats is vital for identifying better methods of exploiting these habitats for vector control. A case in point is the identification of volatile attractant compounds that may emanate from these preferred habitats. These volatile compounds are most likely associated with the nutritional suitability of the habitat to ensure the survivability and fitness of the larval and resulting adult populations. Knowing what aspects of the habitat are used as cues by the adult female to select a suitable oviposition site can provide a novel method for disrupting the life cycle of the mosquito and potentially reducing disease transmission.


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  7. Acknowledgments

We acknowledge the help of Ireneo Briceno, Russell King, and Petr Macek for time and effort in collecting mosquito samples. The help and advice of Doerthe Műller-Navarra and Ann Liston with FA analyses and data evaluation was invaluable. The comments of all the reviewers are highly appreciated. This research was supported by National Institutes of Health Grant R01 AI49726-01.


  1. Top of page
  7. Acknowledgments
  • Ahlgren, G., M. Carlstein, and I.B. Gustafsson. 1999. Effects of natural and commercial diets on the fatty acid content of the European grayling. J. Fish Biol. 55: 11421155.
  • Ahlgren, G., W. Goedkoop, H. Markensten, L. Sonesten, and M. Boberg. 1997. Seasonal variations in food quality for pelagic and benthic invertebrates in Lake Erken – the role of fatty acids. Freshwater Biol. 38: 555570.
  • Arts, T.M., M.T. Brett, and M.J. Kainz (eds.). 2009. Lipids in Aquatic Ecosystems 2nd ed. Springer, Dordrecht .
  • Caramujo, M. J., H.T.S. Boschker, and W. Admiraral. 2008. Fatty acid profiles of algae mark the development and composition of harpacticoid copepods. Freshwater Biol. 53: 7790.
  • Cavaletto, J.F. and W.S. Gradner. 1999. Seasonal dynamics of lipid in freshwater benthic invertebrates. In: M.T. Arts and B.C. Wainman (eds.). Lipids in Freshwater Ecosystems. pp. 109131. Springer- Verlag, New York .
  • Chapman, R.F. 1998. The Insects: Structure and Function. Cambridge University Press, Cambridge .
  • Dadd, R.H. 1981. Essential fatty acids for mosquitoes, other insects and vertebrates. In: G. Bhaskaran, S. Friedman, and J.G. Rodriguez (eds.) Current Topics in Insect Endocrinology and Nutrition. pp. 189214, Plenum, New York .
  • Dadd, R.H. 1983. Essential fatty acids–insects and vertebrates compared. In: Metabolic Aspects of Lipid Nutrition in Insects, T.E. Mittler and R.H. Dadd (eds.) pp. 107147. Westview Press, Boulder , CO .
  • Dadd, R.H. and J.E. Kleinjan. 1979. Essential fatty acid for the mosquito Culex pipiens: arachidonic acid. J. Insect Physiol. 25: 495502.
  • Dadd, R.H., J.E. Kleinjan, and S.M. Asman. 1988. Eicosapentaenoic acid in mosquito tissues: differences between wild and laboratory-reared adults. Environ. Entomol. 17: 172180.
  • Dadd, R.H., J.E. Kleinjan, and V.P. Sneller. 1977. Development of several species of mosquito larvae in fully defined dietary media: preliminary evaluation. Mosq. News 37: 699703.
  • Dadd, R.H., J.E. Kleinjan, and D.W. Stanley-Samuelson. 1987. Polyunsaturated fatty acids of mosquitoes reared with single dietary polyunsaturates. Insect Biochem. 17: 716.
  • Damiens, D., M.Q. Benedict, M. Wille, and J.R.L. Gilles. An inexpensive and effective larval diet for Anopheles arabiensis (Diptera: Culicidae): Eat like a horse, a bird or a fish? J. Med. Entomol. (In Press).
  • Desvilettes, C., G. Bourdier, C. Amblard, and B. Barth. 1997. Use of fatty acids for the assessment of zooplankton grazing on bacteria, protozoans and microalgae. Freshwater Biol. 38: 629637.
  • Gladyshev, M.I., N.N. Sushchik, A.A. Kolmakova, G.S. Kalachova, E.S. Kravchuk, E.A. Ivanova, and O.N. Makhuta. 2007. Seasonal correlations of elemental and omega 3 PUFA composition of seston and dominant phytoplankton species in a eutrophic Siberian Reservoir. Aquat. Ecol. 41: 923.
  • Grieco, J. P., E. Rejmankova, N.L. Achee, C.N. Klein, R. Andre and D. Roberts. 2007. Habitat suitability for three species of Anopheles mosquitoes: Larval growth and survival in reciprocal placement experiments. J. Vector Ecol. 32: 176187.
  • Hayashiya, K. and R.F. Harwood. 1968. Fatty acids of the mosquito Anopheles freebornii. Ann. Entomol. Soc. Am. 61: 278280.
  • Kattner, G. and H.S.G. Fricke. 1986. Simple gas-liquid-chromatographic method for the simultaneous determination of fatty-acids and alcohols in wax esters of marine organisms. J. Chromatog. 361: 263268.
  • Kaufman, M.G., E. Wanja, S. Maknoja, N.M. Bayoh, J. Vulule, and E.D. Walker. 2006. The importance of algal biomass to the growth and development of Anopheles gambiae larvae. J. Med. Entomol. 43: 669676.
  • Kaufman, M.G., E.D. Walker, T.W. Smith, R.W. Merritt, and M.J. Klug. 1999. Effect of larval mosquitoes (Aedes triseriatus) and stemflow on microbial community dynamics in container habitats. Appl, Environ. Microbiol. 65: 26612673.
  • Kuusipalo, L. and R. Käkelä. 2000. Muscle fatty acids as indicators of niche and habitat in Malawian cichlids. Limnol. Oceanogr. 45: 9961000.
  • Manguin, S., D.R. Roberts, R.G. Andre, E. Rejmankova, and S. Hakre, 1996. Characterization of Anopheles darlingi (Diptera: Culicidae) larval habitats in Belize, Central America. J. Med. Entomol. 33: 205211.
  • Marazouzi, C., G. Masson, M.S. Izquierdo, and J.C. Pihan. 2008. Midsummer heat wave effects on lacustrine plankton: Variation of assemblage structure and fatty acid composition. J. Thermal Biol. 33: 287296.
  • Müller-Navarra, D.C. 1995. Biochemical versus mineral limitation in Daphnia. Limnol. Oceanogr. 40: 12091214.
  • Müller-Navarra, D.C. 2008. Food web paradigms: The biochemical view on trophic interactions. Int. Rev. Hydrobiol. 93: 489505.
  • Müller-Navarra, D.C., M.T. Brett, S. Park, S. Chandra, P. Ashley, E.Z. Ballantyne, E. Zorita, and C.R. Goldman. 2004. Unsaturated fatty acid content in seston and tropho-dynamic coupling in lakes. Nature 427: 6972.
  • Müller-Navarra, D.C., M.T. Brett, A.M. Liston, and C.R. Goldman. 2000. A highly unsaturated fatty acid predicts carbon transfer between primary producers and consumers. Nature 403: 7477.
  • Napolitano, G.E. 1999. Fatty acids as trophic and chemical markers in freshwater ecosystems. In: M.T. Arts and B.C. Wainman (eds). Lipids in Freshwater Ecosystems. pp. 2144, Springer – Verlag, New York .
  • Nor Aliza, A.R. and D.W. Stanley. 1998. A digestive phospholipase A2 in larval mosquitoes, Aedes egypti. Insect Biochem. Molec. Biol. 28: 561569.
  • Parrish, C.C. 1999. Determination of total lipid, lipid classes and fatty acids in aquatic samples. In: M.T. Arts and B.C. Wainman (eds). Lipids in Freshwater Ecosystems. pp. 420, Springer – Verlag, New York .
  • Persson, J. and T. Vrede. 2006. Polyunsaturated fatty acids in zooplankton: variation due to taxonomy and trophic position. Freshwater Biol. 51: 887900.
  • Rejmánková, E., D.R. Roberts, R.E. Harbach, E.L. Peyton, S. Manguin, R. Krieg, J. Polanco, and L. Legters. 1993. Environmental and regional determination of Anopheles (Diptera, Culicidae)larval distribution in Belize, Central America. Environ. Entomol. 22: 978992.
  • Rejmánková, E., A. Harbin-Ireland, and M. Lege. 2000. Bacterial abundance in larval habitats of four species of Anopheles (Diptera: Culicidae) in Belize, Central America. J. Vector Ecol. 25: 229238.
  • Rejmánková, E., J. Komárek, and J. Komárková. 2004. Cyanobacteria – a neglected component of biodiversity: patterns of species diversity in inland marshes of northern Belize (Central America). Divers Distrib. 10: 189199.
  • Rejmánková, E., K.O. Pope, D.R. Roberts, M.G. Lege, R. Andre, J. Greico, and Y. Alonzo. 1998. Characterization and detection of Anopheles vestitipennis and Anopheles punctimacula (Diptera: Culicidae) larval habitats in Belize with field survey and SPOT satellite imagery. J. Vector Ecol. 23: 7488.
  • Timmermann, S.E. and H. Briegel. 1996. Effects of plant, fungal and animal diets on mosquito development. Entomol. Exp. Appl. 80: 173176.
  • Walker, E.D. and R.W. Merritt. 1993. Bacterial enrichment in the surface microlayer of Anopheles quadrimaculatus (Diptera:Culicidae) larval habitat. J. Med. Entomol. 30: 10501052.
  • Watton, R.S. and T.M. Preston. 2005. Surface films: areas of water bodies that are often overlooked. BioScience 55: 137145.