Species Diversity and Entomological Inoculation Rate of Anophelines Mosquitoes in an Epidemic Prone Areas of Bure District, Northwestern Ethiopia

doi:10.21203/rs.3.rs-103609/v1

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Species Diversity and Entomological Inoculation Rate of Anophelines Mosquitoes in an Epidemic Prone Areas of Bure District, Northwestern Ethiopia

https://doi.org/10.21203/rs.3.rs-103609/v1

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Background: Malaria is the leading health problem in Ethiopia. The country has been prevented malaria vectors mostly using long-lasting insecticide-treated nets, the application of indoor residual spraying chemicals, and source reductions. Before interventions, identifying the responsible malaria vector in disease transmission (sporozoite rate) is very vital; hence, the present study was designed to assess species diversity and entomological inoculation rate of Anopheles mosquito in Bure district, Northwest Ethiopia.

Methods: Adult mosquitoes were collected from July 2015 to June 2016 using the center for disease control and prevention light traps, pyrethrum spray catches, and artificial pit shelters. Mosquitoes were morphologically identified. Following this, An. gambiae s.l was identified molecularly. Head-thorax sporozoite infectivity of the adult female Anopheles mosquitoes was assessed using enzyme-linked immunosorbent assays.

Results: Morphologically, nine species of the genus Anopheles were identified in the three villages, composed of Anopheles demeilloni, An. arabiensis, An. funestus, An. coustani, An. squamosus, An. cinereus, An. pharoensis, An. rupicolus, and An. natalensis. Of these species, An. demeilloni was the most predominant, whereas An. cinereus, An. rupicolus and An. natalensis were the least representative species (p < 0.0001). Greater number of adult Anopheles mosquitoes were collected in Shnebekuma, non-irrigated villages than non- irrigated village (Workmidr) and irrigated village (Bukta) (p < 0.0001). The overall Plasmodium infective rate (P. falciparum and P. vivax) in the district was 0.31%. The overall annual sporozoite rate in non-irrigated villages (Shnebekuma and Workmidr) was 0.35%, whereas zero in irrigated village (Bukta). The overall estimated EIR of Anopheles mosquitoes was 5.7 infectious bites /person /year for both P. falciparum and P. vivax in the district. The annual EIR Anopheles species in non-irrigated villages was 5.65 ib/p/y, which was higher than irrigated village (0 ib/p/y).

Conclusions: Both the primary (An. arabiensis) and secondary (An. funestus and An. pharoensis) malaria vectors of Ethiopia were identified in the three villages. Three of Anopheles species, An. arabiensis, An. funestus, and An. coustani were found to be infected only in irrigated villages. Source reduction and proper usage of long-lasting insecticide nets and indoor residual spraying could be implemented in the non- irrigated villages to cut the vector abundance and EIR.

Infectious Diseases

An. arabiensis

An. funestus

An. coustani

non-irrigated villages

Bure

Ethiopia

Malaria is transmitted by the blood feeding of infectious female Anopheles mosquitoes [1, 2] with a complex parasite life cycle, which depends on both humans and mosquitoes [3, 4]. In Africa, malaria transmission is varied within districts, between seasons [5, 6, 7], between houses and within agricultural zones [6].

Several factors have reported to influencing the vectorial role of mosquitoes in disease transmission including vector density and heterogeneity, resting and blood-feeding behavior, host preference, longevity, susceptibility to Plasmodium infection, micro-ecological, socioeconomic factors, poor environmental sanitation, and environmental modification [8, 9, 10, 11]. Therefore, a better understanding of these factors is a precondition in planning effective vector control measures [12].

Moreover, EIR is an important indicator of malaria transmission that helps relate both the human-biting activity of the Anopheles vectors and the risk to humans to malaria infection [13, 14]. The dynamics of an EIR are governed by the fluctuation in vector densities and their sporozoite rates [15]; which again influenced by temperature, altitude, rainfall, and urbanization [16].

Therefore, EIR is considered a more direct measure of transmission intensity [16] and its values are used to quantify the impact of vector control measures [17], such as LLINs, IRS, and source reduction [18], malaria endemicity [19] and the risk of an epidemic [20]. Thus, effective vector control measures directly reduce malaria incidence (sporozoite rates) and then EIRs [18, 21, 22].

Generally, understanding the dynamics of malaria transmission in a population is very significant; it provides an insight into the magnitude of the problem and helps to describe when and where the greatest risk occurs. It also helps to facilitate the development and implementation of appropriate control strategies [14, 23, 24].

In Ethiopia, malaria is the leading health problem of the country [25] because three-fourth (75%) of the total area of the country is malarious and more than two-third (approximately 68%) of the total populations live below 2,000 meters above sea level /m.a.s.l/ [26]. In the country, the nature of malaria transmission is seasonal, and unstable [27], vary with elevations, temperature, and rainfall [28, 29].

The major malaria vector in Ethiopia is Anopheles arabiensis while An. pharoensis, An. funestus and An. nili are secondary vectors [30, 31]. The Ethiopian federal ministry of health has four key intervention strategies to prevent and treat malaria: 1) spraying houses with insecticides (IRS), 2) mass distribution of LLINs, 3) having lifesaving drugs and 4) treatments for pregnant women [30].

In particular to Amhara region, Ethiopia, malaria is obviously a serious health problem [32, 33, 34]. Bure district is one of the major malaria focus areas in the region known by the high epidemic, which has claimed many lives [29, 35, 36, 37]. Previously conducted epidemiological studies in the region indicated that the prevalence of malaria has shown a sharp reduction in many districts including Bure surrounding [38]. However, information on entomological indices is incomplete in Amhara region [39] and totally absent in the Bure district in particular. Therefore, the present study was designed to assess species diversity, sporozoite and entomological inoculation rates of Anopheles mosquito in Bure district, Ethiopia.

2.1 Sstudy Area

Longitudinal study was conducted in Bure district, northwestern of Ethiopia, from July 2015 to June 2016. Geographically, Bure district is situated at an altitude ranging from 700 (Blue Nile gorge) to 2,350 m.a.s.l. (Figure 1). Socioeconomically, the majority (85%) of the populations are farmers who grow maize, teff (Eragrostis teff), pepper, potatoes, wheat, millets, followed by beans & peas, sunflower, niger, spices, vegetation’s, and others; and the rests are merchants (6.8%) and others (non-governmental organizations, civil servants) (8.2%). Animals such as cattle, sheep, hens, mules, and donkeys are reared by the farmers. Additionally, both modern and traditional bee-keepers were present. The majority of the populations in the district live in houses made of mud and corrugated iron roofs.

The majority of Bure districts has subtropical zone (Woina-Dega) climate with annual mean minimum and maximum temperature of 9.9°C and 29.2 °C, respectively and 2,000 mm mean annual rainfall range being 1,350 - 2,500 mm. The major rainy season of the district is from July to September, and a small amount is obtained from May to June and from October to December. The rest of the months (January - April) are dry seasons [33].

The study was conducted in three rural villages: Bukta, Workmidr and Shnebekuma, from July 2015 - June 2016. The description of the detail of the three villages are found elsewhere [40]. Totally, these villages are malarious, bed nets were distributed for the three villages once per 3-years before malaria infestation begins, on the first week of September. Moreover, anti-malaria chemical spraying (IRS) (Deltamethrin, K-Othrine Flow) was administered to the three villages according to the national spraying operation guidelines [30].

2.2. Collection, Identification and Processing of Anopheline Mosquitoes

2.2.1. Collection: Anopheles mosquitoes were sampled longitudinally from July 2015 - June 2016. Entomological surveys were conducted monthly in each village, for one year using Center for LTCs, PSCs and APSs. In each village, 9 houses for LTCs and 10 houses for PSCs were randomly selected and scattered in near to the breeding sites, in the middle and periphery sides of the village. In parallel, 27-miniature light traps were prepared to collect the outdoor host seeking mosquitoes for the three villages, each had 9- LTCs. Additionally, six APSs were prepared in three villages to collect outdoor resting mosquitoes; each village had two.

Indoor host-seeking Anopheles mosquitoes were collected from 6:00 PM (sunset) to 6:00 AM (sunrise) by using miniature LTCs (Model 512; J. W. Hock Co., Atlanta, USA) once per month per house [41, 42]. In the same trends, the outdoor host seeking mosquitoes were collected by LTCs from 06:00 AM to 06:00 PM hrs once per month.

Indoor-resting mosquitoes were collected in the mornings from 6:00 AM to 8:30 AM hrs using PSCs for the study period. Collection was made using white floor sheets, hand lenses, Baygon aerosol (Tetramethrin:0.4% and Permethrin:0.4%; SC. Johnson & Son. Inc, USA), small petri-dishes, paper cups with net covers, forceps, cotton wool, and a torch [43]. Additionally, outdoor-resting mosquitoes were collected in the morning from 6:30 AM - 7:30 AM from APSs (1.5 m depths, 1.0 m width, and 1.2 m length) using a handheld mouth aspirator [44]. The number of human occupants and other potential vertebrate hosts in each surveyed house during the previous night were recorded. Moreover, the house condition of each surveyed house was recorded; including the types of house, type of wall, and number of LLINs used and spray status.

Outdoor-resting mosquitoes were collected from artificially made pit shelters by using a handheld mouth aspirator, paper cup with net covers, cotton wool, torch, and pencil. APSs were constructed under the shade of various dense shrub trees 10 - 15 meters away from the resident villages. APSs had 1.5 m depths, 1.0 m width, and 1.2 m length (1.5 m × 1.0 m × 1.2 m). Approximately 0.5 m from the bottom of each pit-shelter, a 30-cm horizontal deep cavity was prepared in each of the four sides [44]. A collection was made from 6:30 AM - 7:30 AM hrs. Before collection begins, the mouth of each pit shelter was covered with insecticide untreated white net to prevent mosquitoes from escaping and for visibility purpose. Resting mosquitoes was collected for about 10 - 20 minutes in each pit.

Identification: Mosquitoes collected by LTCs, PSCs and APSs were identified morphologically at genus level using taxonomic keys [45,46,47]. Culex and male Anopheles were recorded and discarded. Moreover, morphological identified and individually preserved An. gambiae specimens were identified by species-specific PCR [48] at the Molecular Biology Laboratory of Tropical and Infectious Diseases Research Center, Jima University. DNA was extracted from individual preserved An. gambiae complex species based on [49]. Then, DNA amplification was carried out. Following this, gel electrophoresis was carried out [48]. At the end, agarose-gel was placed on UVP (Photo Doc-It- imaging system or UV-Trans- illuminator) to see the nature of the bands (Appendix 1a & b). Those mosquitoes that remained unamplified (without any band on the gel) were tested three times in an independent manner.

2.3 Determination of Plasmodium Infection Rate (SIR) and Entomological Inoculation Rate To determine the infection rate, the heads and thoraces of An. arabiensis, An. funestus group, An. coustani, An. squamosus and An. cinereus females were tested by ELISA for CSP of P. falciparum, P. vivax-210 and P. vivax-247 by the method developed by Wirtz et al. [50]. The dried “head-thorax” portion of individual female Anopheles was placed in 1.5ml eppindorff tube. 50μl of grinding buffer (GB) was added and grinding was made using electrical-pestle. After completion of grinding, the pestle was rinsed with two 100 μl volumes of GB, to have 250 μl final volumes. Then, 50μl of the capture monoclonal antibodies (MAb) of P. falciparum (Cat No: 37-00-24-2; Lot No: 2013pCAP), P. vivax (P.v-210, Lot No: 060415), and P. vivax (P.v-247, Lot No: 030276) were added to labeled 96-wells of separate polyvinyl chloride microtiter plates in duplicates (six in number). And then, the wells covered and incubated at room temperature for 30 minutes. After incubation, well contents were removed, banged five times on a paper towel, and then 200 μl BB was added again in each well, covered, and incubated for one hour to block the remaining active binding sites. Well contents were then aspirated and banged five times on a paper towel, and 50 μl of each mosquito triturate (including positive and negative control) was added according to the prepared sporozoite ELISA-sheets and covered and placed to incubate for two hours. For positive controls, P.f (fragmented stock solution) for P.f-wells, P.v-210 (fragmented stock solution) for P.v-210-wells and P.v-247 (fragmented stock solution) for P.v-244-wells, and for negative control unfed laboratory reared species were used. After 2 hrs of incubation, the mosquito triturates were aspirated and the wells were washed twice with 200 μl PBS-Tween using ELISA-Washer /well washer (Model No: BioTek-WLx 50). Then, 50μl of peroxidase-linked MAbs of P.f (Cat No: 37-00-24-3; Lot No: 2013PfHRR), P.v-210 (Cat No: 37-00-24-3; Lot No: 060414) and P.v-247 (Cat No: 37-00-24-4; Lot No: 030246) were added to respective wells of the microtiter plate, then covered and incubated for one hour. After one hrs incubation, the plates were washed three times with 200 μl PBS-Tween using ELISA-Washer and 100 μl substrate solutions were added in single well and covered and incubated for thirty minutes. Substrate solutions was a mixture of ABTs peroxidase substrate solution-A (Lot No: 080775; Product Code: 50-64-00) and Peroxidase substrate solution-B (Lot No: 080831; Product Code: 50-65-00). Finally, after 30 minutes incubation, absorbance values at 405 nm were recorded from the ELISA plate reader (BioTek-ELx800). Those samples considered positive if the mean absorbance values of the duplicate assays exceeded twice the mean of the three negative controls [50].

2.4 Data Analysis

Data were entered and cleaned using 2007-Microsoft Excel and analyzed using SPSS software package version-20.0 (SPSS, Chicago, IL, USA).

Mean variance between Anopheles and Culex, between the indoor and outdoor host seeking mosquitoes was tested using independent samples T-test (p < 0.05). Variation in mean densities between species and species among villages were analyzed using one-way analysis of variance (ANOVA) (p < 0.05). Significant means (ANOVA) were separated using Tukey test (HSD).

Sporozoite rate for each species was estimated as the number of mosquitoes with sporozoite divided by the number of females examined multiplied by 100 [51]. The annual EIRs were estimated by standard method: 1.605 (No of ELISA positive from CDC light trap/No ELISA tested) X (No of An. arabiensis/ funestus/ coustani collected from CDC light trap/Total No of CDC light traps) x 365 days. Monthly EIR was estimated by a standard method: 1.605 (No. ELISA positive from CDC light trap/Total No. ELISA tested) x (NoAn. arabiensis/ funestus/ coustani collected from CDC light trap/ Total No number of CDC light traps) x No days per month [52]. Before running Independent-samples T-test and ANOVA, normality of data was first checked and transformed [log10(x+1)]. When significant differences were observed in ANOVA, the Tukey test (HSD-Test) was used to separate the means (p < 0.05). All statistical analyses were performed at the 5 % significance level.

3.1 Species Diversity and Abundance of Anopheles Mosquitoes

A total of 11,625 female mosquitoes were collected in Bure district using LTs, PSCs and APSs. Of these, 59.5% (n = 6,922) belonged to the genus Culex while the rest 40.5% (n = 4,703) were from the genus Anopheles. The proportions of the two genera have not shown statistically significant difference (t = 1.165; df = 22; p = 0.257). Morphologically, nine species of the genus Anopheles were identified in the three villages, belonged to Anopheles demeilloni, An. gambiae s.l, An. funestus s.l, An. coustani, An. squamosus, An. cinereus, An. pharoensis, An. rupicolus, and An. natalensis (Fig. 2). Of these species, An. demeilloni was the most predominant whereas An. cinereus, An. rupicolus and An. natalensis were the least representative species (one-ANOVAs: F _{8, 99} = 24.593, p < 0.0001).

Moreover, a total of 66 specimens of An. gambiae s.l was identified to species by PCR, of which 63 (95.5%) were successfully amplified and identified as An. arabiensis (Appendix-1a & b). Only three (4.5%) specimens which were checked for three times were not amplified. Thus, all An. gambiae s.l collected in this study were An. arabiensis.

The greater number of adult Anopheles mosquitoes were collected in Shnebekuma, non-irrigated villages (n = 3,875) than non- irrigated village (Workmidr, n = 472) and irrigated village (Bukta, n = 356) (one ANOVA F_{2, 33} = 19.202, p < 0.0001). The mean density of each Anopheles mosquito by village is presented in Table 1. Significantly higher density of An. arabiensis, An. funestus, An. squamosus and An. demeilloni were recorded from Shnebekuma than other villages (p < 0.05). In Bukta, only An. cinereus was predominant. An. pharoensis, An. rupicolus and An. natalensis were very rare and not distributed uniformly in the three villages; however, the remaining species were abundant in all villages. An. natalensis was recorded only in Shnebekuma village, whereas An. rupicolus was found only in Bukta and Workmidr villages.

Table 1: Mean density of Anopheles species by village, Bure district, Ethiopia (at 3 and 33 degree

of freedom)

Species	Village			F -value	p-value
Species	Bukta	Workmidr	Shnebekuma	F -value	p-value
An. arabiensis	(Mean ± SE) 0.73 ± 0.10^b	(Mean ± SE) 0.79 ± 0.13^b	(Mean ± SE 1.55 ± 0.12^a	16.057	0.0001*
An. pharoensis	0.06 ± 0.05	0.06 ± 0.05	0.15 ± 0.071	0.846	0.438
An. funestus	0.51 ± 0.13^b	0.45 ± 0.11^b	1.48 ± 0.11^a	23.935	0.0001*
An. coustani	0.73 ± 0.16	0.49 ± 0.12	1.11 ± 0.20	2.705	0.082
An. squamosus	0.28 ± 0.11^b	0.27 ± 0.09^b	1.01 ± 0.12^a	15.375	0.0001*
An. cinereus	0.47 ± 0.12^a	0.03 ± 0.03^b	.25 ± 0.09^ab	6.356	0.005*
An. demeilloni	0.60 ± 0.10^b	0.90 ± 0.20^b	2.13 ± 0.10^a	33.832	0.0001*
Note: Mean (s) followed by the same letter (s) in the same row are not significantly different from each other at p < 0.05, Tukey HSD

3.2 Sporozoite Rates of Anophelines Mosquitoes

Annual sporozoite rates of Anopheles mosquito species (collected by using LTCs) are shown in Table 2. In total, 1,474 (1,466 from light traps and 8 from PSCs) Anopheles mosquitoes were tested for Plasmodium Circumsporozoite-Protein (CSPs) which consisted of 488 An. arabiensis, 350 An. funestus, 436 An. coustani, 152 An. squamosus and 40 An. cinereus. Eight specimens from PSCs were excluded for the EIR calculation because they were not infected with sporozoite.

Table 2: Annual sporozoite rates and EIRs of the three Anopheles species in three sites in Bure district, Ethiopia (July 2015 - June 2016)

Villages Name		Sporozoite infected mosquitoes		# Collected		# Tested	# positive for –CSPs			Infective Rate							Annual EIR
										*P.f,* n (%)	**P.v-247, n (%)**		**P.v-210, n (%)**		Total (%)
							CDC-IN	CDC-OUT	Total
Bukta (Irrigated Village)		An. arabiensis		72		56	0	0	0	-	-		-		0		0
Shnebekuma & Workmidr (Non-Irrigated Village)		An. arabiensis		678		432	1	0	0	1(0.23)	-		-		1(0.23)		1.42
Total				750		488	1	0	1	1(0.20)	-		-		1(0.20)		1.4
Bukta (Irrigated Village)		An. funestus		53		20	0	0	0	-	-		-		0		0
Shnebekuma & Workmidr (Non-Irrigated Village)		An. funestus		587		330	0	2	2	-	2(0.61)		-		2(0.61)		3.22
Total				640		350	0	2	2	-	2(0.57)		-		2(0.57)		3.31
Bukta (Irrigated Village)		An. coustani		103		67	0	0	0	-	0		0		0		0
Shnebekuma & Workmidr (Non- Irrigated)		An. coustani		503		369	0	1	1	-	-		1(0.27)		1(0.27)		1.23
Total				606		436	0	1	1	-	-		1(0.23)		1(0.23)		1.3
All species in Irrigated Village (Bukta)				228		143	0	0	0	0	-		0		0		0
All species in Non-Irrigated Villages (Shnebekuma & Workmidr)				1,768		1,131	1	3	4	1(0.09)	2(0.18)		1(0.09)		4(0.35)		5.65
All Total				1,996		1,274	1	3	4	1(0.08)	2(0.16)		1(0.08)		4(0.31)		5.7
Annual Sporozoite Rates and EIRs by Village, Bure district, Ethiopia
Village	Species		Total # of collected species		Total # of Tested Specimens			P.f, n (%)		P.v-247, n (%)		P.v-210, n (%)		Total (%)		Annal EIR
Shenbekuma	An. arabiensis		588		369			1(0.27)		0		0		0.3		2.9
Shenbekuma	An. funestus		544		312			0		1(0.32)		0		0.3		4.3
Shenbekuma	An. coustani		389		297			0		1(0.34)		0		0.3		3.4
Workmidr	An. funestus		34		18			0		0		1(5.6)		5.6		5.1

Of 1,466 Anopheles mosquito specimens examined for CSP, only 4 specimens were positive for CSPs. A single An. arabiensis was positive for P. falciparum and a single An. coustani and two An. funestus specimens were positive for P. vivax. The overall Plasmodium infective rate (P. falciparum and P. vivax) in the district was 0.31% (4/1,274). Infection rate of An. arabiensis, An. funestus, and An. coustani was 0.2% (1/488), 0.57% (2/350) and 0.23% (1/436), respectively. The two infected An. funestus specimens and the single infected An. coustani specimen were collected outdoor while the single An. arabiensis specimen was collected indoor (Table 3).

All sporozoite-infected Anopheles mosquito specimens were collected only from non-irrigated villages. The overall annual sporozoite rate in non-irrigated villages was 0.35%; An. funestus accounted for 0.61% (2/330), An. coustani accounted for 0.27% (1/369), and An. arabiensis accounted for 0.23% (1/432). Of the non-irrigated villages, three infected mosquitoes were found in Shnebekuma (Table 3). Monthly sporozoite rate distribution is shown in Table 3. Infected Anopheles mosquitoes were collected in three different months; July, December, and January.

3.3 Entomological Inoculation Rate of Anopheline Mosquitoes

Estimate annual and monthly EIRs of An. arabiensis, An. funestsu and An. coustani are shown in Tables 2 and 3. The overall estimated EIR of Anopheles mosquitoes was 5.7 infectious bites /person /year (ib/p/y) for both P. falciparum and P. vivax in the district. In particular, the overall P.v EIR of An. funestus, P.f -EIR of An. arabiensis and P.v-EIR of An. coustani were 3.3 ib/p/y, 1.4 ib/p/y and 1.3 ib/p/y, respectively.

Estimates of the EIRs of the three species were also made individually for the three villages (Table 2). The annual EIR of the three Anopheles species in non-irrigated villages was 5.65 ib/p/y, which was higher than irrigated village (0 ib/p/y). This variation was true for each species. The overall P.f-EIR of An. arabiensis was 1.42 ib/p/y in non-irrigated villages and higher than irrigated village (0 ib/p/y). From non-irrigated villages, Shnebekuma had 2.9ib/p/y for An. arabiensis while 0ib/p/y. for Workmidr. The overall P.v-EIR of An. funestus was 3.2 ib/p/y in non–irrigated villages and zero ib/p/y in irrigated village. Of the non-irrigated villages, Workmidr had higher P.v-EIR for An. funestus (5.1 ib/p/y) than Shnebekuma (4.3 ib/p/y). The P.v-EIR of An. coustani was 1.23 ib/p/y in non-irrigated villages. Among non-irrigated villages, only Shnebekuma had 3.4ib/p/y for An. coustani.

Seasonal variation of an EIR is shown in Table 3. The highest monthly EIR for An. arabiensis was 1.6 ib/p/month during the main rainy season; July. Monthly EIR for An. funestus was highest (1.7 ib/p/month) in the dry season (January) followed by a December (1.3 ib/p/month). Monthly EIR for An. coustani, was 1.1 ib/p/month and detected in December.

Table 3

Monthly distribution of CSP–positive (sporozoite rate) and EIR of the three *Anopheles* species using CDC-LTs in Bure district, from July 2015 - June 2016, [July - Sep = Rainy; Oct- Dec and May -June = Small rainy; and Jan - April = Dry Seasons]
Month of collection	Tested Species	# collected specimens	# Tested Specimens	# positive for P.f n (%)	# positive for P.v-247, n (%)	# positive for P.v-210 n (%)	Sporozoite rate n (%)	EIR
July 2015	An. arabiensis	25	14	1(7.14)	0	0	1(7.14%)	1.6
December 2015	An. funestus	16	11	0	1 (9.1)	0	1(9.1)	1.3
January 2016	An. funestus	39	21	0	1(4.8)	0	1(4.8)	1.7
December 2015	An. coustani	44	37	0	0	1(2.7)	1(2.7)	1.1

The overall aim of this study was to examine the diversity and abundance of Anopheles mosquitoes in the three villages. It also helped to determine the sporozoite and entomological inoculation rates of Anopheles mosquitoes in the district. A total of nine Anopheles species belonging to An. arabiensis, An. funestus, An. pharoensis, An. coustani, An. squamosus, An. cinereus, An. demeilloni, An. rupicolus and An. natalensis were recorded in Bure district. An. arabiensis, the main malaria vector in Ethiopia [31, 53, 54], was also among others recorded in such high altitude area (range 2,000–2,157 m.a.s.l), which is in agreement with various findings [55, 56] where An. arabiensis had been recorded at an altitude of 2,110 m.a.s.l and 2,170 m.a.s.l, from the outskirts of Addis Ababa and in Tigray region, respectively. Similarly, many studies that conducted in Ethiopia [57, 58] and in Kenyan highland [59, 60, 61, 62] recorded An. arabiensis at higher altitude.

In our study area, well-known secondary malaria vectors (An. funestus and An. pharoensis) of Ethiopia were recorded. An. funestus was the second most abundant vector which was recorded in the higher altitude. Previously, An. funestus, had been reported from Gojam [63]. However, it is inconsistent with other reports from Ethiopia, where this species had been recorded in areas below 2,000 m.a.s.l [64, 65,66, 67]. The occurrence of this species up to 2,157 m.a.s.l could be attributed to the presence of increased temperature as a result of climate change, land cover and land use changes [62,68,69,70]. Anopheles pharoensis was among the least prevalent species in the study area and it had been reported from high altitude areas (between 2,110 m.a.s.l and 2,200 m.a.s.l) of Ethiopia [55,71]. This species is the secondary malaria vector in Ethiopia below 2,000 m.a.s.l [71, 72,73] though not found infected.

An. coustani was another mosquito recorded at 2,157 m.a.s.l in our study. This observation is in line with various reports [55, 60] that made in the highlands of Ethiopia and Kenya. An. coustani is a known malaria vector in Kenya [61,67,74,75] and was known to be a suspected vector in Ethiopia [62,76,77]. In the present survey, An. coustani was not only abundant (n = 606, 12.9%), but also found infected with Plasmodium vivax. All these information indicated that this species is really would be the other potential malaria vectors in Ethiopia in the future.

In the present study, Anopheles’ mosquitoes were collected in three villages, but highest proportions of adult Anopheles mosquitoes were collected in non-irrigated villages (Shnebekuma and Workmidr) than irrigated village (Bukta). This result was not comparable to other findings that made in Ethiopia [78,79] and in Africa [80, 81, 82].

Higher proportions of mosquito in non-irrigated village could be due to the presence of more productive breeding habitats throughout the study period than irrigated villages [83, Personal obsr.]. On the other hands, the lower densities of Anophelines in irrigated villages (and areas near a dam) are probably due to a greater wealth created in the community via irrigation, which helped to construct good houses, resulted in the prohibition of the mosquitoes to enter in the house; thereby less numbers of mosquitoes were collected. Many studies have approved the purpose of well-constructed houses in reducing the abundances mosquito in the house [71,84,85,86]. In the irrigated village (Bukta), the health center is established at the center of the settlement (inhabitants) than non-irrigated villages. Being very near, the villagers may have a better treatment seeking behavior [6] and health experts may give more attention to irrigated surrounding because it is believed to be malaria is usually common in dammed and irrigated area [72,76,79].

In our study, the overall annual CSP rate of all the Anopheles species (P. falciparum and P. vivax) was 0.31%, which is almost comparable to 0.33% reported from Eritrea [87]. However, it was higher than the other studies from Ethiopia (0.12%) [88] and Tanzania (0.02%) [89]. The highest sporozoite rate in our study is due to the presence of a large proportion of the Anopheles mosquitoes across the year. These populations had better access to human blood meals [8–11]. The expansion of maize farming (maize pollen) in the district is enhanced mosquito populations that can participate directly in cycles of human biting and malaria transmission [90]. On the contrary, the overall sporozoite rate of all Anopheles species in this study is very lower as compared to other reports from different parts of Ethiopia [57,73,91,92,93] and in Africa [82, 94,95]. This very low all over sporozoite rate may be due to the cumulative effects of intensive usage of LLINs and application of IRS, resulted in a decline in human-vector contact, a decrease in the number of mosquitoes and a reduction entomological inoculation rate into people [96–99].

The result of this study showed that the intensity of malaria transmission in the study area is low. In line with the low sporozoite rate, recently made an epidemiological survey from health facilities indicated the reduction of malaria cause in the most Amhara region including Bure-surrounding district [38].

The overall annual P.f-sporozoite rate of An. arabiensis was 0.2%, which is similar to the 0.2% reported from Eritrea [87] and higher than the 0.06% and 0.01% reported from Ethiopia and Eritrea, respectively [88,100]. Various findings reported relatively higher annual sporozoite rates of An. arabiensis in various parts of Ethiopia, 0.3%, 0.28% and 0.3%, respectively [57,92,101]. Similarly, higher annual overall P. f-sporozoite rate of An. arabiensis has been documented in different parts of Ethiopia [66,76] and elsewhere in Africa [82,94,102, 103, 104].

In this study, sporozoite infected An. arabiensis sampled in July as reported by Kibret et al. [73]. In this month, usually malaria transmission occurs in the district (in Amhara region and Ethiopia) [26,105] because a small amount of rain is obtained from May to June which is good for both the vectors and the parasite life cycle.

The overall annual sporozoite rate of An. funestus was 0.57%, is higher as compared to 0.03%, which was reported from Senegal [106]. However, as it is lower as compared with the other reports in Ethiopia [73] and in Africa [7,82,94,104,107,109].

The current study revealed the presence of P. vivax-247 infected An. funestus in the district. However, there was no report showing P. vivax infected An. funestus in Ethiopia until now. P.v-247 infected An. funestus was collected only from outdoor in December and January. Occurrence of an infection in December is consistent with the major malaria transmission season of Amhara region and Ethiopia [26,105].

In this study, An. funestus had a higher sporozoite rate than An. arabiensis and An. coustani. Previously, Fontaine et al.[35] reported that An. gambiae s.l (An. arabiensis) was the only responsible vector for most of the transmissions of malaria in the highland parts of the Amhara region. However, the presence of infectious An. funestus in the current study may suggest that populations of An. funestus could be responsible for the transmission of malaria in the highland fringes of Bure district.

Astonishingly, our study found the third infected species, An. coustani by P. vivax-210. This infectious An. coustani specimen was sampled from outdoor in December. Similar to the present finding, Kelel [110] and Yewhalaw et al. [76] reported the same P. vivax-210 infected An. coustani from Gilgel-Gibe hydroelectric dam area, Ethiopia. The overall annual sporozoite rate for An. coustani was 0.34%, which was lower than other reports in Ethiopia (0.68%) [77] and in Kenya (1.3%) [74]. Likewise, higher sporozoite rate of An. coustani reported from Ethiopia [76] and Kenya [75, 111] than the present study. In Kenya, Taveta district, An. coustani was well known vector of malaria [74]. Moreover, the occurrence of infected An. coustani in December corresponds with the major malaria transmission season in Ethiopia, in particular Bure district [26]. Generally, until now An. coustani was not incriminated as a vector of malaria in Ethiopia, but standing from the current and previous data this species might be a potential malaria vector in Ethiopia. Therefore, it needs a regular entomological surveillance, monitoring and appropriate incrimination of this species to determine its role in malaria transmission in Ethiopia.

The current study revealed the difference in sporozoite rate between villages. All infected mosquitoes were from non-irrigated villages, Shnebekuma and Workmidr. The average annual sporozoite rates of An. arabiensis (0.23%), An. funestus (0.61%) and An. coustani (0.27%) were higher in non-irrigated than irrigated village (nil for all species). Similar to our results, various studies indicated the presence of significantly higher annual sporozoite rate in non-irrigated than irrigated villages in different parts of Africa [5,81,112,113]. Ijumba et al. [112], Appawu et al. [5], and Sogoba et al. [114] also reported highest annual sporozoite rates for An. arabiensis and An. funestus in non-irrigated than irrigated sites (e.g., rice irrigated area) in Africa. The presence of infected mosquitoes in non-irrigated villages might be associated with the occurrence very large numbers of the adult vectors in the two non-irrigated villages (Shnebekuma, n = 3,875; Workmidr, n = 472) than non- irrigated village (Bukta, n = 356). Non-irrigated villages had more breeding habitats throughout the year (Personal obser.), which helped to increase the vector survival rate by releasing enough moisture to the surrounding areas [114]. Therefore, population dynamics and adult age structure are important determining factors for the ability to transmit malaria [115].

However, contradict to the current result, Kibret et al. [116] obtained highest overall annual sporozoite rate of An. arabiensis (1.67%) from the irrigated village than non-irrigated villages (0.43%) in Zeway area, central Ethiopia. Similarly, Jaleta et al. [117] in Ethiopia and Dolo et al. [80], Muturi et al. [81] and Mboera et al. [82] in Africa found highest annual sporozoite rates for An. gambiae s.l and An. funestus in irrigated than non-irrigated sites. The absence of sporozoite infected Anophelines mosquitoes in irrigated village is due to the presence improved housing condition and standard of living because of the wealth created by irrigation. Therefore, mosquitoes are unable to enter into the house and further farmers can protect themselves from nuisance mosquitoes using replaceable health-care facilities [6,80,118]. Health education on malaria is the other means of variation [6,119]. Farmers in irrigated village could have access to malaria information because the health center and primary school was established as the hub of the irrigated village (Personal obser.). Moreover, the absence or the presence of little malaria in irrigated community could be the possible reason for the absence infectious mosquitoes in the irrigated village [80,120].

Generally, this study revealed that P. falciparum and P. vivax were the common Plasmodium species in Bure district. This result was in agreement to the epidemiological study conducted in areas surrounding Bure district and in the other Amhara zones that both P. falciparum and P. vivax were the prevalent parasites [26,38,121, 122].

This study indicated that the combined Entomological inoculation rate for both Anopheles species was 5.7 ib/p/y (overall annual EIR). It was extremely lower than those records from similar altitude and altitudes below this study district. Contrary to our study, high infectivity bites per person per year had been reported in Ethiopia [73,117] and elsewhere in Africa [112,118]. In Chad, Kerah-Hinzoumbe et al. [123] found 311 ib/p/y; in Ghana, Appawu et al. [5] reported 418 ib/p/y and in Malawi Mzilahowa et al. [94] reported 183 ib/p/y.

The estimated EIRs of the three Anopheles mosquito species varied across villages and seasons in Bure district. The estimated overall annual P.fEIR of An. arabiensis was 1.4 ib/p/y. It is very lower as compared with those reported by Antonio-Nkondjio et al. [111] (16.5 ib/p/y), Mboera et al. [82] (728 ib/p/y) and Massebo et al. [101] (17.4 ib/p/y) in Cameroon, Tanzania and southwestern Ethiopia, respectively. In this study, the overall annual P.v EIR of An. funestus was 3.3 ib/p/y. It was lower as compared with other reports [82,111, 107], who reported between 6.5–151.4 ib/p/y, 141ib/p/y and 12ib/p/y in Cameroon, Ghana and Tanzania, respectively. Similarly, An. coustani had lower annual P.vEIR, which was 1.3 ib/p/y. It is not comparable from Antonio-Nkondjio et al. [111] finding, who documented 3.4 ib/p/y in Cameroon.

In our study, infectious mosquitoes were detected only in three months (July, December, and January), though the main malaria vectors were collected in all months. This is highly linked with the given attention (large scale distribution and acceptance of LLINs and the application of IRS) which was offered by the region and the district health offices because all villages in Bure district are malarious. Being this, human-mosquito contact has declined and a reduction sporozoite inoculation rate into people were observed [98,99,124].

A total of nine Anopheles species belonging to An. arabiensis, An. funestus, An. pharoensis, An. coustani, An. squamosus, An. cinereus, An. demeilloni, An. rupicolus and An. natalensis were recorded in Bure district. This finding documented important malaria vectors of Ethiopia (An. arabiensis, An. funestus, An. pharoensis). However, higher proportions of adult Anopheles mosquitoes were collected in non-irrigated villages (Shnebekuma and Workmidr) than irrigated village (Bukta). Anopheles arabiensis, An. funestus and An. coustani found to be infected. The CSP-ELISA results indicated that the prevalence of malaria infectious Anopheles mosquitoes is very low and this makes Bure district suitable for malaria control and elimination. Annual P.f-EIR of An. arabiensis (1.42 ib/p/y), P.v-EIR of An. funestus (3.22 ib/p/y) and P.v-EIR of An. coustani (1.23 ib/p/y) were higher in non-irrigated villages (Shnebekuma and Workmidr) than irrigated (0 ib/p/y) village (Bukta). Even the distribution pattern of the annual EIRs between villages were highly variable, this indicated full attention was only given to irrigated village. Being this, the annual EIR of each species for irrigated village reached zero. Generally, the obtained infective bites per person per year was low (annual EIRs) which is less than ten (EIR < 10); therefore, the finding implied that the study area is considered to have unstable malaria transmission and had a low malaria transmission intensity [126]; however, not < 1, this implied much work remain to be done.

EIR: Entomological Inoculation Rate; LLINs: long-lasting insecticides-treated nets; IRS: indoor residual spraying; LTCs: Disease Control and Prevention Light Trap Catches; PSCs: Pyrethrum Spray Catches; APSs: Artificial Pit- Shelters; ELISA: ELISA: Enzyme-Linked Immunosorbent Assay; CSP: circumsporozoite protein; P.f : Plasmodium falciparum; and P.v = P. vivax.

Acknowledgements

We would like to thank the community of Bukta, Shnebekuma and Workmidr, Bure for their cooperation during our survey time to collect adult mosquitoes. Our deepest gratitude given to Malaria Consortium, Ethiopia, for providing CDC-LTs, full larva collection equipment and internet service. We acknowledge to Dr. Habte Tekie for his enragement and provision of CDC-LT lamps (light-bulbs) and chemicals; Etifanos Kebede, Endalew Zemene and Daneil Emana for supporting lab work and sharing their experiences; Girma Gudesho for preparing study maps; Amanuel T/Mariyam for giving GPS instrument; Dr. Desta Ejeta for providing chemicals. Finally, our appreciate is extended to Addis Ababa and Mizan-Tepi, Universities for providing financial support; Jima University for provision of laboratory facilities.

Competing interests

The authors declare that they have no competing interests.

Authors’ contributions

TA and DY conceived and designed the study; TA conducted field work and experiment in the laboratory, analyzed and interpreted the data, wrote the manuscript; and DY and EG supervised in the field and in laboratory. All authors read and approved the final manuscript.

Funding

This study was financially supported by Addis Ababa, Mizan-Tepi, and Jimma Universities. These universities were not involved in the design of the study, data collection, analysis, and interpretation of data, and in writing the manuscript.

Availability of data and materials

All data presented in this manuscript (study) are available in the hand of the correspondent authors and will release if necessary, with a reasonable request.

Ethics approval and consent to participate

A collection of mosquitoes was carried out after obtaining ethical approval from the ethical review committee of Addis Ababa University (Reference No.: CNSDO/382/07/15), Amhara Health Regional Bureau (Permission Reference No.: H/M/TS/1/350/07) and the Head of the Bure District Health Office (Permission Reference No.: BH/3/519L/2). Moreover, informed consent was obtained from the head of the selected households.

Consent for publication

Not applicable.

Koutsos AC, Blass C, Meister S, Schmidt S, MacCallum RM, Soares MB, et al. Life cycle transcriptome of the malaria mosquito Anopheles gambiae and comparison with the fruit fly Drosophila melanogaster. Natl Acad Sci USA 2007,1- 6.
Sinka ME. Global Distribution of the dominant vector species of malaria. 2013. p. 109- 127.
Garrett-Jones C, Boreham PFL, Pant CP. Feeding habits of anophelines (Diptera: Culicidae) in 1971–1978, with reference to the human blood index: a review. Bull. Entomol Res. 1980, 70: 165-185.
Bannister LH, Sherman IW. Plasmodium. In: Encyclopedia of life sciences (ELS). John Wiley and Sons, Ltd: Chic-heister. 2009. p.1-2.
Appawu MA, Owusu-Agyei S, Dadzie S, Asoala V, Anto F, Koram K, Rogers W, et al. Malaria transmission dynamics at a site in northern Ghana proposed for testing malaria vaccines. Trop Med Int Health. 2004, 9:164-170.
Sissoko MS, Dicko A, Briet OJ, Sissoko M, Sagara I, Keita HD, et al. Malaria incidence in relation to rice cultivation in the irrigated in Sahel of Mali. Acta Tropica 2004, 89: 161-170.
Mboera LE, Mlozi MR, Senkoro KP, Rwegoshora RT, Rumisha SF, Mayal BK, et al. Malaria and agriculture in Tanzania: impact of land-use and agricultural practices on malaria burden in Mvomero district. National Institute for Medical Research, Dares Salaam, Tanzania. 2007.p. 26-34.
Cohuet A, Harris C, Robert V, Fontenille D. Evolutionary forces on Anopheles: What makes a malaria vector? Trends Parasitol 2010, 26:130-136.
Ndoen E, Wild C, Dale P, Sipe N, Dale M. Mosquito longevity, vector capacity, and malaria incidence in west Timor and central Java, Indonesia. Int Scholarly Res Network 2012, 1-5.
Mboera LE, Bwana VM, Rumisha SF, Stanley G, Tungu PK, Malima RC. Spatial abundance and human biting rate of Anopheles arabiensis and An. funestus in savannah and rice agro-ecosystems of central Tanzania. Geospat Health 2015, 10:26-29.
Molina-Cruz A, Zilversmit MM, Neafsey DE, Hart DL, Barillas-Mury C. Mosquito vectors and the globalization of Plasmodium falciparum Malaria. Annu Rev Genet 2016, 50:447-465.
Adeleke AM, Mafinana CF, Idown AB, Samwobo SO, Idown OA. Population dynamics of indoor sampling mosquitoes and their implication in disease transmission in Abeokuta, south-western Nigeria. J Vector Borne Dis 2010, 47:33-38.
Smith T, Charlwood JD, Takken W, Tanner M, Spiegelhalter DJ. Mapping the densities of malaria vectors within a single village. Acta Tropica 1995, 59:1-18.
Smith T, Killeen GF, Lengeler C, Tanner M. Relationships between the outcomes of Plasmodium falciparum infection and the intensity of transmission in Africa. Am J Trop Med Hyg 2004, 71:80-86.
Krafsur ES, Armstrong JC. An integrated view of entomological and parasitological observations on falciparum malaria in Gambela, Western Ethiopian Lowlands. Trans R Soc Trop Med Hyg 1978, 72:348–356.
Obala AA, Kutima HL, Nyamogoba HD, Mwangi AW, Simiyu CJ, Magak GN, et al. Anopheles gambiae and An. arabiensis population densities and infectivity in Kopere village, western Kenya. J Infect Dev Ctries 2012, 6:637-643.
Coosemans M, Wery M, Mouchet J, Carnevale P. Transmission factors in malaria epidemiology and control in Africa. Memorias do Instituto Oswaldo Cruz 1992, 87:385-391.
Shaukat AM, Breman JG, McKenzie FE. Using the entomological inoculation rate to assess the impact of vector control on malaria parasite transmission and elimination. Malar J 2010, 9: 1-8
Burkot TR, Graves PM. The value of vector-based estimates of malaria transmission. Ann Trop Med Parasitol 1995,89:125-134.
Onori E, Grab B. Indicators for the forecasting of malaria epidemics. Bull World Health Organ 1980, 58:91-98.
Smith DL, McKenzie FE. Statics and dynamics of malaria infection in Anopheles mosquitoes. Malar J 2004, 3:1-12.
Killeen GF, Smith TA. Exploring the contributions of bed nets, cattle, insecticides and excito- repellency to malaria control: a deterministic model of mosquito host seeking behaviour and mortality. Trans R Soc Trop Med Hyg 2007, 101: 867-880.
Smith DL, Dushoff J, Snow RW, Hay SI. The entomological inoculation rate and Plasmodium falciparum infection in African children. Nature 2005, 438:492-8.
Hay SI, Smith D, Snow R. Measuring malaria endemicity from intense to interrupted transmission. Lancet Infect Dis 2008, 8:369-378.
The Carter Center. Summary proceedings 4th annual malaria control program review Ethiopia and Nigeria. Carter Center, Atlanta, Georgia; 2013. P.1-19.
Ayele DG, Zewotir TT, Mwambi HG. Prevalence and risk factors of malaria in Ethiopia. Malar J 2012, 11:1-8.
Ministry of Health, MoH. National Strategic Plan for Malaria Prevention Control and Elimination in Ethiopia 2011-2015. Ethiopia Ministry of Health. Addis Ababa, Ethiopia; 2010.
Graves PM, Richards FO, Ngondi J, Emerson PM, Shargie EB, Endeshaw T, et al. Individual, household and environmental risk factors for malaria infection in Amhara, Oromia and SNNP regions of Ethiopia. Trans R Soc Trop Med Hyg 2009, 103:1211-1220.
Connor SJ, Dinku T, Wolde-Georgis T, Bekele E, Jima D. A collaborate epidemic early warning and response initiative in Ethiopia. 2010. p. 1-4.
MoH. National Malaria Guidelines. (3rd edt.). Addis Ababa, Ethiopia. Ethiopia Ministry of Health. Addis Ababa, Ethiopia; 2012.
The President's malaria initiative, PMI. Eleventh annual report to congress; 2017.
Tamiru MA, Kassa AW, Beyene BB, Mossie TB, Mekonnen YA. Malaria out-break investigation in Mecha, Dera and Fogera districts, Amhara region, Ethiopia. Am J Health Res 2014, 2:182-187.
Midekisa A, Beyene B, Mihretie A, Bayabil E, Wimberly MC. Seasonal associations of climatic drivers and malaria in the highlands of Ethiopia. Parasit Vectors 2015, 8:1-11.
Lake MW, Mebratur M, Mehari D, Dessie K. Epidemiological analysis of malaria outbreak in Ankesha district, Awi zone, Amhara region, Ethiopia, 2012: Weaknesses in control measures and risk factors. Sci J Public Health 2016, 4:132-137.
Fontaine RE, Najjar AE, Prince JS. The 1958 malaria epidemic in Ethiopia. Am J Trop Med Hyg 1961, 10:795 - 803.
Kiszewski A, Teklehaimanot A. A review of the clinical and epidemiologic burdens of epidemic malaria. Am J Trop Med Hyg 2004,71: 128-135.
Kassa AW, Tamiru MA, Yeshanew AZ. Assessment of control measure and trends of malaria in Burie-Zuria district, west Gojjam zone, Amhara region, north west Ethiopia. Malar Res Treatment 2015,1-4.
Toyama Y, Ota M, Molla G, Beyene BB. Sharp decline of malaria cases in the Burie-Zuria, Dembia, and Mecha districts, Amhara region, Ethiopia, 2012–2014: Descriptive analysis of surveillance data. Malar J 2016, 15:1-8.
Vajda EA, Webb CE. Assessing the risk factors associated with malaria in the highlands of Ethiopia: What do we need to know? Trop Med Infect Dis 2017, 2:1-13.
Adugna T, Getu E, Yewhalaw D. Species diversity and distribution of Anopheles mosquitoes in Bure district, Northwestern Ethiopia. Heliyon 2020, 6: e05063.gna.
Lines JD, Curtis CF, Wilkes TJ, Njunwa KJ. Monitoring human-biting mosquitoes (Diptera: Culicidae) in Tanzania with light-traps hung beside mosquito. Bull Entomol Res 1991, 81: 77-84.
Mboera LE, Kihonda J, Braks MA, Knols BG. Influence of centers for disease control light trap position, relative to a human-baited bed-net, on catches of Anopheles gambiae and Culex quinquefasciatus in Tanzania. Am J Trop Med Hyg 1998, 59: 595-596.
World Health Organization, WHO. Malaria Entomology and Vector Control: Learner’s Guide. (Trial eds.). World Health Organization HIV/AIDS, Tuberculosis and Malaria, Roll Back Malaria. HO/CDS/ CPE/SMT/2002.18. Rev.1: Part I; 2003.
WHO. WHO manual on practical entomology in malaria, Part-II, WHO, Geneva;1975.
Verrone GA. Outline for the determination of malaria mosquitoes in Ethiopia: Part- I- Adult female Anopheles. Mosq New 1962, 22: 37-50
Gillies MT, Coetzee MA supplement to the Anophelinae of Africa South of the Sahara (Afrotropical Region). Johannesburg. South Afr Insect Med Research 1987, 55:1-125.
Glick JI. Illustrated kit to the female Anopheles of southwestern Asia and Egypt (Diptera: Culicide). Mosq Systematic 1992, 24:125-151.
Wilkins EE, Howell PI, Benedict MQ. IMP PCR primers detect single nucleotide poly- morphisms for Anopheles gambiae species identification, Mopti and Sahavanna rDNA types, and resistance to dieldrin in Anopheles Arabiensis. Malar J. 2006, 5:1-7.
DNeasy^®Blood and Tissue Kits. DNeasy Blood and Tissue Handbook. www.qiagen.com.handbooks. Accessed 1 Jan 2011.
Wirtz RA. Sporozoites ELISA direction. Centers for Disease Control and Prevention (CDC), Atlanta, Georgia, USA; 2016.
WHO. Malaria entomology and vector control: Guide for participants. NLM classification: WC 765). Ingram Publishing. Geneva, Switzerland; 2013.
Drakeley C, Schellenberg D, Kihonda J, Sousa CA, Arez AP, Lopes D, et al. An estimation of the entomological inoculation rate for Ifakara: a semi-urban area in a region of intense malaria transmission in Tanzania. Trop Med Int Health 2003,8:767-774.
White GB, Tesfaye F, Boreham PF, Lemma G. Malaria vector capacity of Anopheles arabiensis and Anopheles quadriannulatus in Ethiopia: chromosomal interpretation after six years storage of field preparations. Trans R Soc Trop Med Hyg 1980,74: 683-684.
Balkew M, Elhassen I, Ibrahim M, Gebre-Michael T, Engers H. Very high DDT-resistant population of Anopheles pharoensis Theobald (Diptera: Culicidae) from Gorgora, northern Ethiopia. Parasite 2006, 3:327- 329.
Woyessa A, Gebre-Micheal T, Ali A. An indigenous malaria transmission in the outskirts of Addis Ababa, Akaki town and its environs. Ethiop J Health 2004, 81:2-7.
Dejenie T, Yohannes M, Assmelash T. Adult mosquito populations and their health impact around and far from dams in Tigray Region, Ethiopia. Ethiop J Health Sci 2012, 4:40-51.
Animut A, Balkew M, Gebre-Michael T, Lindtjorn B. Blood meal sources and entomological inoculation rates of Anophelines along a highland altitudinal transect in south central Ethiopia. Malar J 2013a, 12:112-121.
Tesfaye S, Belyhun Y, Teklu T, Mengesha T, Petros B. Malaria prevalence pattern observed in the highland fringe of Butajira, Southern Ethiopia: a longitudinal study from parasitological and entomological survey. Malar J 2011, 10:1-9.
Ndenga B, Githeko A, Omukunda E, Munyekenye G, Atieli H, Wamai P, et al. Population dynamics of malaria vectors in western Kenya highlands. J Med Entomol 2006, 43:200- 206.
Chrispinus SM, Moses MN, Donald NS, John MV. Diversity of Anopheles and prevalence of malaria in a highland area of Western Kenya. J Parasitol Vector Biol 2011,3: 33-39.
Mulambalah CS, Siamba DN, Ngeiywa MM, Vulule JM. Anopheles species diversity and breeding habitat distribution and the prospect for focused malaria control in the western highlands of Kenya. Int J Trop Med 2011, 6:44-51.
Kweka EJ, Munga S, Himeidan Y, Githeko AK, Yan G. Assessment of mosquito larval productivity among different land use types for targeted malaria vector control in the western Kenya highlands. Parasit Vectors 2015, 8:1-6.
O’Connor CT. The distribution of anopheline mosquitoes in Ethiopia. Mosq News 1967, 27: 42-55.
Gone T, Balkew T, Gebre-Michael T. Comparative entomological study on ecology and behaviour of Anopheles mosquitoes in highland and lowland localities of Derashe district, southern Ethiopia. Parasit Vectors 2014; 7:1-9.
Ototo EN, Mbugi JP, Wanjala CL, Zhou G, Githeko AK, Yan G. Surveillance of malaria vector population density and biting behaviour in western Kenya. Malar J, 2015, 14:1-9.
Daygena TY, Massebo F, Lindtjorn B. Variation in species composition and infection rates of Anopheles mosquitoes at different altitudinal transects, and the risk of malaria in the highland of Dirashe Woreda, south Ethiopia. Parasit Vectors 2017, 10:1-11.
Degefa T, Yewhalaw D, Zhou G, Lee M, Atieli H, Githeko AK, Yan G. Indoor and outdoor malaria vector surveillance in western Kenya: implications for better understanding of residual transmission. Malar J 2017,16: 1-12.
Hay SI, Rogers DJ, Toomer JF, Snow RW. Annual Plasmodium falciparum entomological inoculation rates (EIR) across Africa: literature survey, internet access and review. Trans R Soc Trop Med Hyg 2000, 94:113-127.
Afrane YA, Zhou G, Lawson BW, Githeko AK, Yan G. Effects of microclimatic changes caused by deforestation on the survivorship and reproductive fitness of Anopheles gambiae in western Kenya highlands. Am J Trop Med Hyg 2006, 74:772-778.
Afrane YA, Githeko AK, Yan G. The ecology of Anopheles mosquitoes under climate change: case studies from the effects of environmental changes in east Africa highlands. Ann N Y Acad Sci 2012, 1249:1-7.
Animut A, Balkew M, Lindtjorn B. Impact of housing condition on indoor biting and indoor- resting Anopheles arabiensis density in a highland area, central Ethiopia. Malar J 2013b, 12:123-130.
Kibret S, Petros B, Boelee E, Tekie H. Entomological studies on the impact of a small-scale irrigation scheme on malaria transmission around Zeway, Ethiopia. Addis Ababa. 2008. p 418-435
Kibret S, Wilson GG, Ryder D, Tekie H, Petros B. Malaria impact of large dams at different eco-epidemiological settings in Ethiopia. Trop Med Health 2017, 45:1-14.
Mwangangi JM, Mbogo CM, Orindi BO, Muturi EJ, Midega JT, Nzovu J, et al. Shifts in malaria vector species composition and transmission dynamics along the Kenyan coast over the past 20- years. Malar J 2013, 12:1-9.
Ogola E, Villinger J, Mabuka D, Omondi D, Orindi B, Mutunga J, et al. Composition of Anopheles mosquitoes, their blood-meal hosts, & Plasmodium falciparum infection rates in three islands with disparate bed net coverage in Lake Victoria, Kenya. Malar J 2017, 16:1-11
Yewhalaw D, Kelel M, Getu E, Temam S, Wessel G. Blood meal sources and sporozoite rates of Anophelines in Gilgel-Gibe dam area, Southwestern Ethiopia. African J Vectors 2014; xxx: xxx- xxx. https: www.researchgate.net/publication/292615632_Blood_meal_so urces_and_sporozoite_rates_of_Anophelines_in_Gilgel-Gibe_dam_area_Southwestern_Et hiopia. Accessed 20 Sep 2020.
Degefa T, Zeynudin A, Godesso A, Haile-Michael Y, Eba K, Zemene E, et al. Malaria incidence and assessment of entomological indices among resettled communities in Ethiopia: a longitudinal study. Malar J 2015, 14:1-10.
Kibret S, McCartney M, Lautze J, Jayasinghe NM. Malaria transmission in the vicinity of impounded water: evidence from the Koka Reservoir, Ethiopia. Colombo, Sri Lanka: international water management institute (IWMI) research report 132, Colombo Sri Lanka. 2009 p. 1-34
Kibret S, Alemu Y, Boelee E, Tekie T, Alemu D, Petros B. The impact of a small-scale irrigation scheme on malaria transmission in Ziway area, Central Ethiopia. Trop Med Int Health 2010, 15:41–50.
Dolo G, Briet OJ, Dao A, Traore SF, Bouare M, Sogoba N, et al. Malaria transmission in relation to rice cultivation in the irrigated Sahel of Mali. Acta Tropica 2004, 89:147-159.
Muturi EJ, Muriu S, Shililu J, Mwangangi S, Jacob BG, Mbogo C, Githure J, Novak RJ. Effect of rice cultivation on malaria transmission in central Kenya. Am Trop Med Hyg 2008, 78: 270-275.
Mboera LE, Senkoro KP, Mayala BK, Rumisha SF, Rwegoshora RT, Mlozi MR, Shayo EH. Spatio-temporal variation in malaria transmission intensity in five agro-ecosystem in Mvomero district Tanzania. Geopat Health 2010, 4:167-178.
Mwangangi JM, Midega J, Kahindi S, Njoroge L, Nzovu J, Githure J, et al. Mosquito species abundance and diversity in Malindi, Kenya and their potential implication in pathogen transmission. Parasitol Res 2012, 110:61-71.
Konradsen F, Amerasinghe P, Der Hoek W, Amerasinghe F, Perera D, Piyaratne M. Strong association between house characteristics and malaria vectors in Sri Lanka. Am J Trop Med Hyg 2003; 68:177-181.
Atieli H, Menya D, Githeko A, Scott A. House design modifications reduce indoor resting malaria vector densities in rice irrigation scheme area in western Kenya. Malar J 2009, 8: 1-9.
Njie M, Dilger E, Lindsay SW, Kirby MJ. Importance of eaves to house entry by anopheline, but not culicine mosquitoes. J Med Entomol 2009, 46:505-510.
Shililu J, Ghebremeskel T, Seulu F, Mengistu S, Fekadu S, Zerom M, et al. Larva habitat diversity and ecology of anopheline larvae in Eritrea. J Med Entomol 2003, 40: 921–929.
Getachew D, Gebre‑Michae T, Balkew, M, Tekie, H. Species composition, blood meal hosts and Plasmodium infection rates of Anopheles mosquitoes in Ghibe River Basin, southwestern Ethiopia. Parasit Vectors 2019, 12:257.
Mwanziva CE, Kitau J, Tungu PK, Mweya CN, Mkali H, Ndege CM, et al. Transmission intensity and malaria vector population structure in Magugu, Babati District in northern Tanzania. J Health Res 201,13: 68-71.
Kebede A, Mccann JC, Kiszewski AE, Ye-Ebiyo Y. New evidence of the effects of agro- ecologic change on malaria transmission. Am J Trop Med Hyg 2005, 73: 676- 680.
Taye A, Haddis M, Adugna N, Tilahun D, Wirtz, A. Biting behavior and Plasmodium infection rates of Anopheles arabiensis from Sille, Ethiopia. Acta Tropica 2006, 97: 50-54.
Tirados I, Costntni C, Gibson G, Torr SJ. Blood-feeding behaviour of the malarial mosquito Anopheles arabiensis: implications for vector control. Med Vet Entomol 2006, 20:425- 437.
Abraham M, Massebo F, Lindtjorn B. High entomological inoculation rate of malaria vectors in area of high coverage of interventions in southwest Ethiopia: Implication for residual malaria transmission. Parasit Epidemiology Control 2017, 2: 61- 69.
Mzilahowa T, Hastings IM, Molyneux ME, McCall PJ. Entomological indices of malaria transmission in Chikhwawa district, Southern Malawi. Malar J 2012, 11:1-9.
Lwetoijera DW, Harris C, Kiware SS, Dongus S, Devine GJ, McCall, PJ, Majambere S. Increasing role of Anopheles funestus and Anopheles arabiensis in malaria transmission in the Kilombero Valley, Tanzania. Malar J 2014, 13:1-10.
Norris LC, Fornadel CM, Hung W, Pineda FG, Norris DE. Frequency of multiple blood meals taken in a single gonotrophic cycle by Anopheles arabiensis mosquitoes in Macha, Zambia. Am J Trop Med Hyg 2010, 83:33-37.
Jima D, Getachew A, Bilak H, Steketee RS, Emerson PM & Graves PM, et al. Malaria indicator survey 2007, Ethiopia: coverage and use of major malaria prevention and control interventions. Malar J 2010, 9:1-11.
Bekele D, Belyhun Y, Petros B, Deressa W. Assessment of the effect of insecticide treated nets and indoor residual spraying for malaria control in three rural kebeles of Adami Tulu district, south central Ethiopia. Malar J 2012, 11:1-9.98.
Bekele D, Petros B, Deressa W, Belyhun Y. Decline of malaria using combined lasting inseti- cide treated nets and DDT house spraying strategies in Adami Tulu Jido Kombolcha District, Central Ethiopia: A Longitudinal Study from parasitologica and entomological indices data. 2013, 2:1-8.
Waka M, Hopkins RJ, Akinpelu O, Curtis C. Transmission of malaria in the Tesseney area of Eritrea: parasite prevalence in children, and vector density, host preference, and sporozoite rate. J Vector Ecol 2005, 30: 27-32.
Massebo F, Balkew M, Gebre-Michael T, Lindtjorn B. Entomological inoculation rates of Anopheles arabiensis in southwestern Ethiopia. Am J Trop Med Hyg 2013, 89:466- 473.
Appawu MA, Bosompem KM, Dadzie S, McKakpo US, Anim-Baidoo I, Dykstra E, et al. Detection of malaria sporozoites by standard ELISA and VecTest^TM dipstick assay in field-collected Anopheline mosquitoes from a malaria endemic site in Ghana. Trop Med Int Health 2003, 8:1012-1017.
Shililu J, Ghebremeskel T, Seulu F, Mengistu S, Fekadu S, Zerom M, et al. Seasonal abundance, vector behavior, and malaria parasite transmission in Eritrea. J Am Mosq Control Assoc 2004, 20:155-164.
Cohuet A, Simard F, Wondji CS, Antonio-Nkondjio C, Awono-Ambene P, Fontenille, D. High malaria transmission intensity due to Anopheles funestus (Diptera: Culicidae) in a village of savannah-forest transition area in Cameroon. J Med Entomol 2004, 41: 901-905.
Midekisa A, Senay G, Henebry GM, Semuniguse P, Wimberly MC. Remote sensing-based time series models for malaria early warning in the highlands of Ethiopia. Malar J 2012, 11:1-9.
Dia I, Konate L, Samb B, Sarr J, Diop, A., Rogerie, F, et al. Bionomics of malaria vectors and relationship with malaria transmission and epidemiology in three physiographic zones in the Senegal River Basin. Acta Tropica 2008, 105:145-153.
Dery DB, Brown C, Asante KP, Adams M, Dosoo D, Amenga-Etego S, et al. Patterns and seasonality of malaria transmission in the forests avannah transitional zones of Ghana. Malar J 2010, 9:1-8.
McCann RS, Ochomo E, Bayoh MN, Vulule JM, Hamel MJ, Gimnig JE, et al. Reemergence of Anopheles funestus as a vector of Plasmodium falciparum in western Kenya after long-term implementation of insecticide-treated bed nets. Am J Trop Med Hyg 2014, 90: 597-604.
Stevenson JC, Pinchoff J, Muleba M, Lupiya J, Chilusu H, Mwelwa I, et al. Spatio-tempora heterogeneity of malaria vectors in northern Zambia: implications for vector control. Parasit Vectors 2016; 9:1-3.
Kelel M. Anopheles mosquitoes host preference and malaria transmission intensity using immunological diagnostic methods in Gilgel-gibe dam area, southwestern Ethiopia. Addis Ababa University. [M.s.c. Thesis]. 2010. p. 37- 41.
Antonio-Nkondjio C, Kerah CH, Simard F, Awono-Ambene P, Chouaibou M, Tchuinkam T, et al. Complexity of the malaria vectorial system in Cameroon: contribution of secondary vectors to malaria transmission. J Med Entomol 2006, 43:1215 -1221.
Ijumba JN, Mosha F, Lindsay S. Malaria transmission risk variation derived from different agricultural practices in an irrigated area northern Tanzania. Med Vet Entomol 2002, 16: 28-38.
Diakite NR, Guindo‑Coulibaly N, Adja AM, Ouattara M, Coulibaly JT, Utzinger J, et al. Spatial and temporal variation of malaria entomological parameters at the onset of a hydro- agricultural development in central Côte d’Ivoire. Malar J 2015, 14:1- 10.
Sogoba N, Doumbia S, Vounatsou P, Bagayok MM, Dolo G, Traore SF, et al. Malaria transmission dynamics in Niono, Mali: The effect of the irrigation systems. Acta Tropica 2007, 101:232-240.
Beck-Johnson LM, Nelson WA, Paaijmans KP, Read AF, Thomas MB, Bjornstad ON. The effect of temperature on Anopheles mosquito population dynamics and the potential for malaria transmission. PLoS ONE 2013, 8:1-10
Kibret S, Wilson GG, Tekie H, Petros, P. Increased malaria transmission around irrigation schemes in Ethiopia and the potential of canal water management for malaria vector control. Malar J 2014; 13:360.
Jaleta KT, Hill SR, Seyoum E, Balkew M, Gebre-Michael T, Ignell R, et al. Agro-ecosystems impact malaria Prevalence: large-scale irrigation drives vector population in western Ethiopia. Malar J 2013, 12:1-11.
Diuk-Wasser MA, Toure MB, Dolo G, Bagayoko M, Sogoba N, Traore SF, et al. Vector abundance and malaria transmission in rice-Grow in villages in Mali. Am J Trop Med Hyg 2005, 72: 725-731.
Audibert M, Josseran R., Josse R, Adjidji A. Irrigation, schistosomiasis and malaria in the Logone Valley, Cameroon. Am J Trop Med Hyg 1990, 42: 550-560.
Ijumba JN, Lindsay SW. Impact of irrigation on malaria in Africa: paddies. Med Vet Entomol 2001, 15:1-11.
Emerson PM, Ngondi J, Biru E, Graves PM, Ejigsemahu Y, Gebre T, et al. Integrating an NTD with one of ‘‘the big three’’: combined malaria and trachoma survey in Amhara region of Ethiopia. PLoS Negl Trop Dis 2008, 2:1-10.
Alemu A, Muluye D, Mihret M, Adugna M, Gebeyaw M. Ten years trend analysis of malaria prevalence in Kola Diba, North Gondar, Northwest Ethiopia. Parasit Vectors 2012, 5:1-6.
Kerah-Hinzoumbe C, Peka M, AntonioNkondjio C, Donan-Gouni1 I, Awono-Ambene P, Same- Ekobo, A. et. al. Malaria vectors and transmission dynamics in Goulmoun, a rural city in south-western Chad. BMC Infect Dis 2009, 9:1-10.
Magesa SM, Wilkes TJ, Mnzava AE, Njunwa K, Myamba J, Kivuyo MD, et al. Trial of pyrethroid impregnated bed nets in an area of Tanzania holoendemic for malaria Part-2. Effects on the malaria vector population. Acta Trop 1991, 49: 97-108.
Takken W, Lindsay SW. Factors affecting the vectorial competence of Anopheles gambiae: a question of scale. Wageningen University. 2003. p.78- 85.

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Species Diversity and Entomological Inoculation Rate of Anophelines Mosquitoes in an Epidemic Prone Areas of Bure District, Northwestern Ethiopia

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Abstract

Figures

1. Introduction

2. Materials And Methods

2.1 Sstudy Area

3. Results

3.1 Species Diversity and Abundance of Anopheles Mosquitoes

3.2 Sporozoite Rates of Anophelines Mosquitoes

3.3 Entomological Inoculation Rate of Anopheline Mosquitoes

4. Discussions

Conclusions

Abbreviations

Declarations

References

Supplementary Files

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