Interactions Between Glossina Pallidipes Salivary Gland Hypertrophy Virus and Tsetse Endosymbionts in Wild Tsetse Populations

doi:10.21203/rs.3.rs-1139411/v1

Download PDF

Research Article

Interactions Between Glossina Pallidipes Salivary Gland Hypertrophy Virus and Tsetse Endosymbionts in Wild Tsetse Populations

https://doi.org/10.21203/rs.3.rs-1139411/v1

This work is licensed under a CC BY 4.0 License

Version 1

posted

You are reading this older preprint version

Read the latest preprint version →

Background

Therefore, tsetse control is considered an effective and sustainable tactic for the control of cyclically transmitted trypanosomosis in the absence of effective vaccines and inexpensive, effective drugs. The sterile insect technique (SIT) is currently used to eliminate tsetse fly populations in an area-wide integrated pest management (AW-IPM) context in Senegal. For SIT, tsetse mass-rearing is a major milestone that associated microbes can influence. Tsetse flies can be infected with micro-organisms, including the primary and obligate Wigglesworthia glossinidia, the commensal Sodalis glossinidius, and Wolbachia pipientis. In addition, tsetse populations often carry a pathogenic DNA virus, the Glossina pallidipes Salivary Gland Hypertrophy Virus (GpSGHV) that hinders tsetse fertility and fecundity. Interactions between symbionts and pathogens might affect the performance of the insect host.

Methods

In the present study, we assessed the possible interaction of GpSGHV and tsetse endosymbionts under field conditions to decipher the bidirectional interactions in different Glossina species. We determined the co-infection pattern of GpSGHV and Wolbachia in natural tsetse populations. We further analyzed the interaction of both Wolbachia and GpSGHV infection with Sodalis and Wigglesworthia density using qPCR.

Results

The results indicated that the co-infection of GpSGHV and Wolbachia was most prevalent in Glossina austeni and Glossina morsitans morsitans, with an explicit significant negative correlation between GpSGHV and Wigglesworthia infection. GpSGHV infection levels of more than 10⁴ were not observed when Wolbachia infection was present at high density (>10^8.5), suggesting a potential protective role of Wolbachia against GpSGHV.

Conclusion

The result indicates that Wolbachia infection might protect tsetse fly against GpSGHV and the interactions between the tsetse host and its associated microbes are dynamic, likely species-specific and significant differences may exist between laboratory and field conditions.

Parasitology

Hytrosaviridae

tsetse microbiota

virus transmission

Wigglesworthia

Wolbachia

Sodalis

Mutualistic bacteria are functionally essential to the physiological well-being of their animal hosts. They benefit their hosts by providing essential nutrients, aiding in digestion and maintaining intestinal equilibrium. Furthermore, mutualistic symbionts foster the development, differentiation, and proper function of their host’s immune system [1–5]. Insects provide a useful model for studying host-microbe interactions because they are associated with bacterial communities that can be easily manipulated during their host’s development [6]. Tsetse flies (Glossina spp.) accommodate various types of bacteria, including two gut-associated bacterial symbionts, the obligate Wigglesworthia glossinidia, and the commensal Sodalis glossinidius, the widespread symbiont Wolbachia pipientis, and the recently discovered Spiroplasma endosymbiont [7–13]. In addition, tsetse flies can house different types of viral infection, including the salivary gland hypertrophy virus (GpSGHV), iflavirus, and negevirus, besides trypanosome parasites [14–17]. Symbiotic associations between insect disease vectors and gut and endosymbiotic bacteria have been particularly well studied to determine how these microbes influence their host’s ability to be infected and transmit disease [18–22]. For example, in tsetse flies, the obligate bacteria Wigglesworthia glossinidia are essential for maintaining the female fecundity and the host immune system through providing an important nutritional component (vitamin B6), but also produce folate (vitamin B9), which aids the trypanosome infection in tsetse making tsetse flies with this obligate symbiont a more suspectable and competent vector of the disease [22–24]. In addition, Sodalis may modulate tsetse’s susceptibility to infection with trypanosomes and several studies using field-captured tsetse have noted that the prevalence of trypanosome infections positively correlates with increased Sodalis density in the fly’s gut [25–29]. In contrast, the exogenous bacterium Kosakonia cowanii inhibits trypanosome infection by creating an unfavorable environment for trypanosome establishment in the mid-gut [30].

Flies in the genus Glossina (tsetse flies) are unique to Africa and are of great medical and economic importance as they serve as a vector for the trypanosomes, the responsible agent for sleeping sickness in humans (human African trypanosomosis or HAT) and nagana in animals (African animal trypanosomosis or AAT) [31, 32]. The presence of tsetse and trypanosomes is considered one of the major challenges for sustainable development in Africa [33, 34]. The lack of adequate and affordable vaccines coupled with pathogen resistance to drug treatments severely limits AAT control, leaving vector control as the most feasible option for sustainable management of the disease [31, 32]. In addition to various pesticide- and trapping-based methods for tsetse control, the sterile insect technique (SIT) is considered, when implemented in the frame of area-wide integrated pest management (AW-IPM) as an efficient, sustainable and environmentally-friendly method [35, 36]. However, the SIT requires the mass-rearing of a large number of males to be sterilized with ionizing-radiation before release into the targeted area [33, 37].

Tsetse fly biology is characterized by its viviparous reproduction rendering tsetse mass-rearing a real challenge. Tsetse flies can nourish intrauterine offspring from glandular secretions and give birth to fully developed larvae (obligate adenotrophic viviparity) [38, 39]. They also live considerably longer than other vector insects, which somewhat compensates for their slow reproduction rate [40]. The ability to nourish larvae on the milk gland secretion, although limiting the number of larvae produced (8-12) per female lifetime, helps to maternally transmit endosymbiont bacteria and pathogens from females to larvae such as Wigglesworthia, Sodalis, Wolbachia, Spiroplasma, and GpSGHV [8, 10, 13, 41]. Moreover, as strictly hematophagous, tsetse relies on the associated endosymbionts to obtain essential elements for female reproduction. Therefore, the well-being of tsetse mass-rearing for SIT is affected by the status of its endosymbionts as well as the infection with pathogenic viruses and the interaction between them. Although Wigglesworthia is an obligate endosymbiont and found in all tsetse species, Sodalis, Wolbachia, and Spiroplasma infection varied from one species to another [8, 10, 42–45]. In addition, infection with the GpSGHV, although reported in different tsetse species, is symptomatic mainly in G. pallidipes [46–48].

The variable responses of different tsetse species to the GpSGHV infection might indicate a possibility of the tsetse microbiota modulating the molecular dialogue between the virus, the symbiont, and the host, shaping the response of each species to the virus infection. It was necessary, therefore, to investigate the infection status of the major tsetse endosymbionts (Wigglesworthia, Sodalis, and Wolbachia) in different tsetse species and their potential interaction. We have recently investigated the interaction between GpSGHV and tsetse symbionts in six tsetse species after virus injection under laboratory conditions [49]. The results indicated that the interaction between the GpSGHV and tsetse symbionts is a complicated process that varied from one tsetse species to another. It is worth noting that the study of Demirbas-Uzel et al. [49] was conducted in tsetse flies maintained under laboratory, controlled conditions (sustainable food availability, constant environmental conditions (temperature and humidity), and high density of the flies), which favors the increase of tsetse symbionts [45, 50, 50–52]. In addition, this study was done using adults artificially infected with GpSGHV by injection.

As the above conclusions may not reflect what happens under natural conditions, in the present study, we aimed to investigate the interaction of the GpSGHV and tsetse symbionts in field-collected samples by evaluating the prevalence of co-infection of GpSGHV and Wolbachia and their interaction with Wigglesworthia and Sodalis infection in natural tsetse populations and comparing the results with those reported under laboratory conditions. The results are also discussed in the context of developing an effective and robust mass production system of high-quality sterile tsetse flies for implementing SIT programs.

Tsetse samples, extraction of total DNA, and PCR amplifications

The field collection of tsetse fly samples, DNA extraction, as well as the PCR-based prevalence of GpSGHV and Wolbachia infections were reported previously [7, 47, 53, 54]. Based on these publications, and using G. m. morsitans, G. pallidipes, G. medicorum, G. brevipalpis and G. austeni samples collected from Burkina Faso, South Africa, Tanzania, Zambia and Zimbabwe four groups of infection patterns were determined: A: flies infected with both GpSGHV and Wolbachia (W⁺/V⁺), B: flies infected only Wolbachia alone (W⁺/V⁻), C: flies infected only with GpSGHV (W⁻/V⁺) and D: flies without GpSGHV or Wolbachia infection (W⁻/V⁻).

Analysis of the interaction between SGHV and Wolbachia, Sodalis, and Wigglesworthia infection in wild tsetse populations

The interaction between GpSGHV and Wolbachia, Sodalis, and Wigglesworthia was assessed by qPCR analysis. Tsetse fly samples were selected for qPCR analysis only if a given population of each species was characterized by the presence of two or three infection groups (W⁺/V⁺), (W⁺/V⁻), and (W⁻/V⁺). Based on this criterion, 203 individual flies (78, 103, and 22 flies with infection status of (W⁺/V⁺), (W⁺/V⁻) and (W⁻/V⁺), respectively) were analyzed (Table 1). The qPCR analysis was performed as previously described [47, 55, 56]. In brief, total DNA was diluted 10-fold before being used for qPCR analysis on a CFX96 real-time PCR detection system (Bio-Rad, Hercules, CA) using the primers and conditions presented in Supplementary Table 1. qPCR data was corrected through the multiplication by the inverse dilution factor to reflect the GpSGHV, Wolbachia, Wigglesworthia or Sodalis copy number (hereafter mention as density) per fly. Analysis of the Wolbachia, Wigglesworthia, Sodalis, and SGHV infection levels was based only on qPCR data with the expected melting curves at 85.5-86°C, 78.5-80°C, 81.5-82°C, and 76.5-77°C, respectively. Data with a melting curve outside the indicated range were excluded from the analysis. The status of Sodalis and Wigglesworthia infection of the samples used for the qPCR analysis was not determined by traditional PCR.

Table 1

SGHV and *Wolbachia* infection status of tsetse flies in natural populations of different *Glossina* species.
Glossina taxon	Country (Area, Collection Date)	N	W+/V+	W+/V-	W-/V+	W-/V-	χ²	P
G. austeni	Tanzania (Jozani, 1997) ¹	42	0	22	2	18
G. austeni	Tanzania (Zanzibar, 1995) ^1*	78	3	72	0	3
G. austeni	South Africa (Zululand, 1999) ^1*	83	51	28	1	3
G. austeni	Coastal Tanzania (Muhoro, NA)	2	0	2	0	0
G. austeni	All locations	205	54	124	3	24	4.32	0.04
G. brevipalpis	South Africa (Zululand, 1995) ¹	50	0	1	0	49
G. brevipalpis	Coastal Tanzania (Muhoro, NA)	1	0	1	0	0
G. brevipalpis	Coastal Tanzania (Muyuyu, NA)	1	0	1	0	0
G. brevipalpis	All locations	52	0	3	0	49
G. f. fuscipes	Uganda (Buvuma island, 1994) ^{1, 2}	53	0	0	6	47
G. medicorum	Burkina Faso (Comoe, 2008)^*	94	2	18	7	67	0.01	0.94
G. m. submorsitans	Burkina Faso (Nazinga, 2009)	3	0	0	0	3
G. m. submorsitans	Burkina Faso (Comoe Folonzo, 2007)	30	0	2	3	25
G. m. submorsitans	Burkina Faso (Comoe, 2008)^*	109	0	4	9	96
G. m. submorsitans	All locations	142	0	6	12	124	0.58	0.45
G. p. palpalis	Democratic Republic of Congo (Zaire, 1995) ¹	48	0	0	1	47
G. tachinoides	Burkina Faso (Nazinga, 2009)	15	0	0	0	15
G. tachinoides	Burkina Faso (Comoe Folonzo, 2007)	112	3	2	26	81
G. tachinoides	Burkina Faso (Comoe, 2008)	72	0	0	8	64
G. tachinoides	Ghana (Pong Tamale, Walewale, 2008)	46	0	5	0	41
G. tachinoides	Ghana (Walewale, 2008)	149	0	27	6	116
G. tachinoides	Ghana (Fumbissi, 2008)	39	0	0	0	39
G. tachinoides	All locations	433	3	34	40	356	0.15	0.70
G. m. morsitans	Coastal Tanzania (Utete, NA)	3	0	2	0	1
G. m. morsitans	Zambia (MFWE, Eastern Zambia, 2007) ^1*	122	26	96	0	0
G. m. morsitans	Tanzania (Ruma, 2005) ^1*	100	29	71	0	0
G. m. morsitans	Zimbabwe (Gokwe, 2006) ¹	74	0	7	8	59
G. m. morsitans	Zimbabwe (Kemukura, 2006) ¹	26	0	26	0	0
G. m. morsitans	Zimbabwe (M.Chiuy, 1994) ^1*	36	5	28	0	3
G. m. morsitans	Zimbabwe (Makuti, 2006) ^1*	99	11	84	1	3
G. m. morsitans	Zimbabwe (Mukond, 1994) ¹	36	0	35	0	1
G. m. morsitans	Zimbabwe (Mushumb, 2006) ¹	8	0	3	0	5
G. m. morsitans	Zimbabwe (Rukomeshi, 2006) ^1*	100	8	90	0	2
G. m. morsitans	All locations	604	79	442	9	74	1.07	0.30
G. pallidipes	Zambia (MFWE, Eastern Zambia, 2007)^1*	203	1	4	97	101
G. pallidipes	Kenya (Mewa, Katotoi, Meru national park, 2007) ¹	470	0	0	10	460
G. pallidipes	Ethiopia (Arba Minch, 2007) ¹	454	0	2	87	365
G. pallidipes	Tanzania (Ruma, 2005) ^1*	83	2	1	42	38
G. pallidipes	Tanzania (Mlembuli and Tunguli, 2009) ¹	94	0	0	0	94
G. pallidipes	Zimbabwe (Mushumb, 2006) ¹	50	0	0	1	49
G. pallidipes	Zimbabwe (Gokwe, 2006) ¹	150	0	0	19	131
G. pallidipes	Zimbabwe (Rukomeshi, 2006) ¹	59	0	5	0	54
G. pallidipes	Zimbabwe (Makuti, 2006) ^1*	96	1	3	5	87
G. pallidipes	Mainland Tanzania (Death Valley, NA)	6	0	4	0	2
G. pallidipes	Coastal Tanzania (Muhoro, NA)	4	0	3	0	1
G. pallidipes	Coastal Tanzania (Muyuyu, NA)	3	0	3	0	0
G. pallidipes	All locations	1672	4	25	261	1382	0.09	0.76
G. p. gambiensis	Senegal (Diacksao Peul and Pout, 2009) ¹	188	0	1	31	156
G. p. gambiensis	Guinea (Kansaba, Mini Pontda, Kindoya, Ghada Oundou, 2009)¹	180	0	0	13	167
G. p. gambiensis	Guinea (Alahine, 2009) ¹	29	0	0	3	26
G. p. gambiensis	Guinea (Boureya Kolonko, 2009) ¹	36	0	0	1	35
G. p. gambiensis	Guinea (Fefe, 2009) ¹	29	0	0	1	28
G. p. gambiensis	Guinea (Kansaba, 2009) ¹	19	0	0	4	15
G. p. gambiensis	Guinea (Kindoya, 2009) ¹	12	0	1	0	11
G. p. gambiensis	Guinea (Lemonako, 2009) ¹	30	0	0	4	26
G. p. gambiensis	Guinea (Togoue, 2009) ¹	32	0	0	1	31
G. p. gambiensis	Guinea (Conakry, 2010)	138	0	5	0	133
G. p. gambiensis	Burkina Faso (Comoe, 2008)	12	0	0	7	5
G. p. gambiensis	Burkina Faso (Comoe Folonzo, 2007)	53	0	1	14	38
G. p. gambiensis	Burkina Faso (Kenedougou, 2007)	37	0	1	0	36
G. p. gambiensis	Burkina Faso (Houet Bama, 2007)	69	0	1	41	27
G. p. gambiensis	Guinea (Fefe, Togoue, Alahine, Boureya Kolonko, 2009-2010)	94	0	5	0	89
G. p. gambiensis	Guinea (Boureya Kolonko, Kansaba, Kindoya, Ghada Oundou, 2009-2010)	94	0	3	0	91
G. p. gambiensis	Mali (Fijira, 2009)	14	0	0	0	14
G. p. gambiensis	Senegal (Diaka Madia, 2009)	42	0	0	0	42
G. p. gambiensis	Senegal (Tambacounda, 2008)	38	0	3	0	35
G. p. gambiensis	Senegal (Simenti, 2008)	33	0	6	0	27
G. p. gambiensis	Senegal (Kédougou, 2008)	15	0	1	0	14
G. p. gambiensis	All locations	1194	0	28	120	1046	3.20	0.07
¹: In these samples, the presence of Wolbachia was tested in Doudoumis et al. 2012.
²: The individuals of G. f. fuscipes were considered negative for Wolbachia based on the results of the initial PCR amplification. The results from the reamplification method were not taken into account so that the conditions were consistent for all species.
^: Samples used for qPCR analysis to determine the density of Wigglesworthia, Sodalis, Wolbachia* and GpSGHV

Statistical analysis

The proportion of single and double infections (GpSGHV and Wolbachia) in wild flies was analyzed by location, species and for all samples together using the Chi-squared test. The Chi-squared tests for independence, Spearman correlation coefficient, and Cochran–Mantel–Haenszel test for repeated tests of independence were performed using Excel 2010. The P-values were calculated from the data with the significance threshold selected as 0.05.

The difference in Wigglesworthia, Sodalis, Wolbachia, and GpSGHV density between different location and tsetse species and the correlation between densities as well as preparing figures were executed in R v 4.0.5 [57] using RStudio v 1.4.1106 [58, 59] with packages ggplot2 v3.3.2.1 [60], lattice v0.20-41 [61], car [62], ggthems [63] and MASS v7.3-51.6 [64]. All regression analyses of symbionts and GpSGHV densities were conducted using the generalized linear model (glm) for different tsetse species and different countries with analysis of deviance Table (Type II tests). The analysis details are presented in Supplementary file 1. Overall similarities in Wolbachia, Wigglesworthia, Sodalis and GpSGHV density levels between tsetse species, countries and infection status were shown using the matrix display and metric multidimensional scaling (mMDS) plot with bootstrap averages in PRIMER version 7+ and were displayed with a Bray and Curtis matrix based on the square root transformation [65]. The tests were based on the multivariate null hypothesis via the non-parametric statistical method PERMANOVA [66]. The Permanova test was conducted on the average of the qPCR density data based on the country-species-sample.

Prevalence of coinfection with GpSGHV and Wolbachia in wild tsetse flies

Analysis of the Wolbachia and GpSGHV infection status for each individual tsetse adult in the previously reported data [7,47,53,54] indicated that the single infection rate was 10.21% (n = 459) and 15.12% (n = 680) for GpSGHV and Wolbachia, respectively, overall taxa and locations combined (Supplementary Figure 1A). No Wolbachia infection was found in two taxa, G. f. fuscipes and G. p. palpalis, and these were excluded from further examination; each was represented by only a single collection (Table 1). A Cochran–Mantel–Haenszel test for repeated tests of independence showed that infection with GpSGHV and Wolbachia did not deviate from independence across all taxa (χ²_MH = 0.848, df = 1, n.s.) and individual Chi-squared tests for independence for each taxon did not show any significant deviation from independence at the Bonferroni corrected α = 0.00714 (Supplementary Table 2). The prevalence of co-infection of GpSGHV and Wolbachia (W⁺/V⁺) in wild tsetse populations varied based on the taxon and the location (Table 1 and Supplementary Figure 1B). No co-infection was found in G. brevipalpis, G. m. submorsitans, and G. p. gambiensis and co-infection was absent in many locations in the remaining taxa. However, a low prevalence of co-infection was found in G. medicorum (2%), G. tachinoides (0.7%) and G. pallidipes (0.2%). A relatively high prevalence of co-infection was only observed in G. austeni (26%), and G. m. morsitans (13%) (Supplementary Figure 1B).

Impact of co-infection (W⁺/V⁺) on GpSGHV, Wolbachia, Sodalis and Wigglesworthia infection density

GpSGHV density

The GpSGHV qPCR data showed overall no statistically significant difference between flies with different infection status (W⁺/V⁺), (W^-/V⁺) and (W⁺/V^-) (X² =1.4625, df = 2, P = 0.481) regardless of tsetse taxon (Supplementary figure 2A). Moreover, no significant difference in GpSGHV copy number was observed between tsetse taxa (X² = 0.752, df = 3, P = 0.861) (Figure 1A). However, a significant difference in the virus copy number was observed between different countries (X² = 16.234, df = 4, P = 0.0027) where the virus copy number in the flies collected from Zambia was significantly lower than those collected from South Africa, Tanzania, and Zimbabwe (Supplementary figure 3A, Supplementary file 1).

Wolbachia density

The copy number of Wolbachia infection was significantly different between tsetse taxa (X² = 6.568, df = 2, P = 0.037) (Figure 1B), and between the infection status (X² = 23.723, df = 2, P << 0.001) (Supplementary figure 2B) and between the countries (X² = 73.507, df = 3, P << 0.001) (Supplementary figure 3B). Wolbachia density was significantly higher in G. m. morsitans than in G. austeni (t = 2.029, P = 0.0478). (Figure 1B, Supplementary file 1).

Overall, a significant difference in Wolbachia density was observed in the flies with different infection status previously determined by conventional PCR, and flies with a (W⁺/V^-) infection pattern showed significantly higher Wolbachia density than flies with a (W⁺/V⁺) infection pattern regardless of the tsetse species (X² = 23.723, df = 2, P << 0.001). This trend was observed in G. m. morsitans (t = 3.184, P = 0.0022) (Supplementary file 1). The Wolbachia density was highest in the flies collected from Zambia (Supplementary figure 3B). Analyzing only the flies with co-infection (W⁺/V⁺) indicated that the Wolbachia density was statistically significantly higher in G. m. morsitans than in G. austeni (t = -2.353, P = 0.024), (Supplementary figure 4B, Supplementary file 1).

Interaction between GpSGHV infection and Wolbachia, Wigglesworthia, and Sodalis infection

The qPCR results of both Wigglesworthia and Sodalis in tsetse adults with different infection status (W⁺/V⁺), (W^-/V⁺) and (W⁺/V^-) indicated that Wigglesworthia density varies significantly between different infection status (X² = 10.706, df = 2, P = 0.0047) and its density in flies with co-infection (W⁺/V⁺) was significantly lower than those with Wolbachia infection only (W⁺/V^-) (t = 3.137, P = 0.0024) but did not differ significantly from flies with virus infection only (W^-/V⁺) (t= 1.656, P = 0.102) (Supplementary figure 2C). Wigglesworthia density varies also between tsetse taxa (X² = 33.479, df = 4, P << 0.001) with higher density in G. m. morsitans and G. pallidipes than G. austeni (Figure 1C) as well as between countries (X² = 19.785, df = 3, P < 0.001) (Supplementary figure 3C, Supplementary file 1).

Sodalis density also varies between tsetse taxa (X² = 21.612, df = 3, P < 0.001) (Figure 1D) and between countries (X² = 21.179, df = 4, P < 0.001) (Supplementary figure 3D) but there was no significant difference between tsetse flies with different infection status (X² = 0.63888, df = 2, P = 0.727) (Supplementary figure 2D, Supplementary file 1).

Analyzing the pairwise correlation between the GpSGHV and each of the tsetse endosymbionts in G. austeni and G. m. morsitans (species with the highest number of flies with coinfection) indicated that the GpSGHV has a significant negative correlation with Wolbachia infection (r = -0.371, t = -2.821, df = 50, P = 0.007) (Figure 2). This trend was observed only with G. m. morsitans (r = -0.474, t = -3.321, df = 38, P = 0.002) but not in G. austeni where the trend was not significant (r = 0.370, t = 1.259, df = 10, P = 0.237) (Supplementary file 1). No flies were observed with high virus density (> 10⁴ copy number) when Wolbachia density was high (~10^8.5 copy number). Contrary to Wolbachia, GpSGHV has significant positive correlation with Wigglesworthia (r = 0.443, t = 3.495, df = 50, P = 0.001) but no correlation with Sodalis infection (r = 0.240, t = 1.749, df = 50, P = 0.086). Wolbachia infection also showed significant negative correlation with Wigglesworthia infection (r = -0.428, t = -3.3496, df = 50, P = 0.001). This trend was only observed in G. m. morsitans (r = -0.637, t = -5.0948, df = 38, P << 0.001) but not in G. austeni (r = 0.119, t = 0.38071, df = 10, P = 0.711). In addition, no significant correlation was found between Wolbachia and Sodalis densities (r= 0.259, t = 1.9007, df = 50, P = 0.063). No flies with high Wigglesworthia density (~10⁸ copy number) were detected when Wolbachia density was high (>10⁸ copy number). Sodalis and Wigglesworthia infections were not correlated (r = 0.257, t = 1.885, df = 50, P = 0.065) (Figure 2, Supplementary file 1).

The qPCR results showed that Wolbachia infected flies had a relatively high Wolbachia density (~10⁷ copies/fly) compared to the GpSGHV and other tsetse symbionts (Wigglesworthia and Sodalis) regardless of the species, country, or infection status (Figure 3). The heat map analysis of the qPCR data of G. austeni and G. m. morsitans clearly indicates the contrast between Wolbachia copy number and Wigglesworthia copy number taking into consideration the infection status, the tsetse taxa, or the countries. In addition, it clearly shows the low copy number of GpSGHV in the samples showing a high Wolbachia copy number (Figure 3, Supplementary figure 5). The bootstrap averages of the metric multidimensional scaling (mMDS) produced clusters based on the species, the country, and the infection status (Figure 4). The Permanova analysis of the density of the GpSGH, Wolbachia, Wigglesworthia and Sodalis based on the country, tsetse species and infection status indicated that the clusters observed between infection status (P = 0.026) and country (P = 0.001) were statistically significant. The interaction between country and infection status was not statistically significant (P = 0.123) (Table 2).

Table 2. PERMANOVA table of results for country and infection status factors and their combinations

Source	df	SS	MS	Pseudo-F	P(perm)	Uniqueperms
Country	2	657.53	328.77	13.003	0.001	999
Species	0	0		No test
Infection status	2	188.97	94.487	3.7369	0.026	998
Country x Species	0	0		No test
Country x Infection status	2	100.59	50.296	1.9892	0.123	999
Species x Infection status	0	0		No test
Country x Species x Infection status	0	0		No test
Res	104	2629.6	25.285
Total	113	4681.3

Within the table, statistically significant differences (P < 0.05) are shown in bold. Perm(s) = permutations.

The screening for GpSGHV and Wolbachia co-infection in natural tsetse populations clearly indicated that the infections in the tested tsetse species were independent. However, when it occurs, it seems that the virus infection might be controlled by Wolbachia infection as demonstrated by the absence of tsetse adults with a high density of GpSGHV in the presence of Wolbachia with high density. Moreover, the screen of wild tsetse populations for GpSGHV and Wolbachia infection indicated that not all Glossina species harbor Wolbachia or GpSGHV. Furthermore, Wolbachia and GpSGHV prevalence was found to differ not only between different tsetse host species but also between different populations within the same tsetse species [7, 11, 47, 53, 54, 67].

The negative impact of Wolbachia infection on the GpSGHV infection in natural tsetse populations is in agreement with the recent report on the interaction of Wolbachia and GpSGHV infection in colonized tsetse populations [49]. However, it is important to note that the setting of this study have major differences from the design of the study of Demirbas-Uzel et al. (2021). These differences include that in our study the fly sex, age, and feeding status were not recorded. In addition, our results present a snapshot of the virus and symbiont infection status at the collection time of the tsetse flies used in the present study, which might suffer from lack of food, different environmental conditions (temperature and humidity), and trypanosome infections. In addition, the number of tested flies was not equally distributed between the tsetse taxa and location. This might explain the lack of co-infection in some taxa and the low number of taxa (G. austeni and G. m. morsitans) used for investigating the interactions between the GpSGHV and tsetse symbionts. The negative correlation between Wolbachia infection and GpSGHV infection was also reported in wild-caught G. f. fuscipes collected from Uganda [67]. This conflicts with our findings as no G. f. fuscipes flies with GpSGHV were reported, which might be due to the low number of tested flies used in our study (n = 53).

Several reports have discussed and well documented the negative effect of Wolbachia on RNA viruses in different insect models such as mosquitoes and Drosophila [68–70], although there have also been reports about Wolbachia enhancement of both RNA and DNA viruses [71, 72]. It is worth mentioning that the negative impact of Wolbachia on GpSGHV was observed only when Wolbachia density was high as the results show the absence of high density (>10⁴) GpSGHV infection with high-density Wolbachia infection (>10^8.5). However, at low Wolbachia density co-infection occurs with a prevalence of more than 10%. The negative impact of high-density Wolbachia infection on GpSGHV is in agreement with previous reports suggesting that the negative impact of Wolbachia on insect viruses is density-dependent [73, 74].

The assessment of the infection density (copy number per fly) of all four microbes (GpSGHV, Wolbachia, Wigglesworthia, and Sodalis) in the same tsetse flies indicated that Wolbachia infection at high density has a significant negative impact on Wigglesworthia infection in G. m. morsitans but not in G. austeni. However, the latter might be due to the low number of analyzed G. austeni flies (n = 21) compared to G. m. morsitans (n = 91). On the other hand, Wolbachia density levels do not correlate with Sodalis. The nature of the negative interaction between Wolbachia and Wigglesworthia is unclear. Whether this negative correlation between Wolbachia and Wigglesworthia is present in other tsetse species beyond G. m. morsitans remains to be seen.

The positive correlation between GpSGHV infection and Wigglesworthia infection observed in G. m. morsitans conflicts with the negative correlation observed in the same species of colonized flies [49]. This result might reflect a specific adaptation between a specific strain of Wigglesworthia which reacts in a specific way to increase its density in the presence of GpSGHV as a manner to restore and enhance the host immune system against the virus infection [75]. The difference in the interaction between the GpSGHV and Wigglesworthia between the results of this study and the results of Demirbas-Uzel et al. [49] might be due to: (i) difference in the host strain/genotype as the tsetse flies of G. m. morsitans were collected from several countries in east Africa (Tanzania, Zambia and Zimbabwe) while the colonized flies originated from Zimbabwe and maintained in the colony since 1997; (ii) different strain(s) of Wigglesworthia circulating in the field samples compared to the ones present in colonized flies [56]; (iii) different strain(s) of the GpSGHV in the field samples [53] and (iv) the difference between field and laboratory conditions whereas the stress from handling the large number of flies in high density in the laboratory might negatively affect Wigglesworthia density levels and/or performance. The same reasons may also explain the difference observed between field and laboratory samples in respect to the interactions between GpSGHV and Sodalis.

Taken together, the present study, despite its limitations in respect to the size of samples and the lack of knowledge about the age, the nutritional and trypanosome infection status as well as the environmental conditions at the time of collection of field specimens, clearly indicates that the interactions between the tsetse host and its associated microbes are dynamic, likely species-specific and significant differences may exist between laboratory and field conditions.

Data availability

Materials described in the paper, including all relevant raw data, are available in this link

https://dataverse.harvard.edu/dataset.xhtml?persistentId=doi:10.7910/DVN/X15PQF

Author Contributions

MMD and DUG: performed the experiments, analyzed the data, and drafted the manuscript. AAA and VD: performed the experiments, and critically revised the manuscript. AGP: analyzed data and critically revised the manuscript. GT: critically revised the manuscript. KB: conceived the study, designed the experiments, interpreted the data, contributed to the drafting, and critically revised the manuscript. AMMA conceived the study, designed the experiments, interpreted the data, and drafted the manuscript. All authors approved the final version of the manuscript and agreed to be accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved.

Funding

This study was supported by the Joint FAO/IAEA Insect Pest Control Subprogramme.

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

The authors would like to thank Ms Carmen Marin, Mr Adun Henry and Mr Abdul Hasim. Mohammed for their technical support.

Chu H, Mazmanian SK. Innate immune recognition of the microbiota promotes host-microbial symbiosis. Nat Immunol. 2013;14:668–75.
Khosravi A, Yáñez A, Price JG, Chow A, Merad M, Goodridge HS, et al. Gut microbiota promote hematopoiesis to control bacterial infection. Cell Host Microbe. 2014;15:374–81.
Douglas AE. Multiorganismal insects: diversity and function of resident microorganisms. Annu Rev Entomol. 2015;60:17–34.
Gomez de Agüero M, Ganal-Vonarburg SC, Fuhrer T, Rupp S, Uchimura Y, Li H, et al. The maternal microbiota drives early postnatal innate immune development. Science. 2016;351:1296–302.
Marchesi JR, Adams DH, Fava F, Hermes GDA, Hirschfield GM, Hold G, et al. The gut microbiota and host health: a new clinical frontier. Gut. 2016;65:330–9.
Benoit JB, Vigneron A, Broderick NA, Wu Y, Sun JS, Carlson JR, et al. Symbiont-induced odorant binding proteins mediate insect host hematopoiesis. eLife. 2017;6:e19535.
Doudoumis V, Tsiamis G, Wamwiri F, Brelsfoard C, Alam U, Aksoy E, et al. Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse flies (genus Glossina). BMC Micobiology [Internet]. 2012;12:S3-. Available from: doi:10.1186/1471-2180-12-S1-S3
Doudoumis V, Blow F, Saridaki A, Augustinos AA, Dyer NA, Goodhead IB, et al. Challenging the Wigglesworthia, Sodalis, Wolbachia symbiosis dogma in tsetse flies: Spiroplasma is present in both laboratory and natural populations. Sci Rep. 2017;7:4699.
Maltz MA, Weiss BL, O’Neill M, Wu Y, Aksoy S. OmpA-mediated biofilm formation is essential for commensal bacterium Sodalis glossinidius to colonize the tsetse fly gut. Appl Environ Microbiol [Internet]. 2012;78:7760–8. Available from: http://dx.doi.org/10.1128/AEM.01858-12
Wang J, Brelsfoard C, Wu Y, Aksoy S. Intercommunity effects on microbiome and GpSGHV density regulation in tsetse flies. J Invertebr Pathol [Internet]. 2013;112:S32–9. Available from: DOI: 10.1016/j.jip.2012.03.028
Wang J, Weiss BL, Aksoy S. Tsetse fly microbiota: form and function. Front Cell Infect Microbiol [Internet]. Frontiers; 2013 [cited 2020 Aug 3];3. Available from: https://www.frontiersin.org/articles/10.3389/fcimb.2013.00069/full
Schneider DI, Saarman N, Onyango MG, Hyseni C, Opiro R, Echodu R, et al. Spatio-temporal distribution of Spiroplasma infections in the tsetse fly (Glossina fuscipes fuscipes) in northern Uganda. Caljon G, editor. PLoS Negl Trop Dis [Internet]. 2019 [cited 2021 Mar 8];13:e0007340. Available from: https://dx.plos.org/10.1371/journal.pntd.0007340
Son JH, Weiss BL, Schneider DI, Dera K-SM, Gstöttenmayer F, Opiro R, et al. Infection with endosymbiotic Spiroplasma disrupts tsetse (Glossina fuscipes fuscipes) metabolic and reproductive homeostasis. PLoS Pathog. 2021;17:e1009539.
Abd-Alla AMM, Cousserans F, Parker AG, Jehle JA, Parker NJ, Vlak JM, et al. Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. J Virol [Internet]. 2008;82:4595–611. Available from: doi:10.1128/JVI.02588-07
Abd-Alla AM, Kariithi HM, Cousserans F, Parker NJ, Ince IA, Scully ED, et al. Comprehensive annotation of the Glossina pallidipes salivary gland hypertrophy virus from Ethiopian tsetse flies: A proteogenomics approach. J Gen Virol. 2016;97:1010–31.
Demirbas-Uzel G, Kariithi HM, Parker AG, Vreysen MJB, Mach RL, Abd-Alla AMM. Susceptibility of tsetse species to Glossina pallidipes salivary gland hypertrophy virus (GpSGHV). Front Microbiol [Internet]. 2018 [cited 2018 Apr 17];9:701. Available from: https://www.frontiersin.org/articles/10.3389/fmicb.2018.00701/full
Meki I, Huditz H-I, Strunov A, Van Der Vlugt R, Kariithi HM, Rezaezapanah M, et al. Characterization and tissue tropism of newly identified iflavirus and negevirus in tsetse flies Glossina morsitans morsitans<\i>. 2021. 2021;In Press.
Weiss B, Aksoy S. Microbiome influences on insect host vector competence. Trends Parasitol [Internet]. 2011;27:514–22. Available from: doi:10.1016/j.pt.2011.05.001
Narasimhan S, Fikrig E. Tick microbiome: the force within. Trends Parasitol. 2015;31:315–23.
Dey R, Joshi AB, Oliveira F, Pereira L, Guimarães-Costa AB, Serafim TD, et al. Gut Microbes Egested during Bites of Infected Sand Flies Augment Severity of Leishmaniasis via Inflammasome-Derived IL-1β. Cell Host Microbe. 2018;23:134-143.e6.
Song X, Wang M, Dong L, Zhu H, Wang J. PGRP-LD mediates A. stephensi vector competency by regulating homeostasis of microbiota-induced peritrophic matrix synthesis. PLoS Pathog. 2018;14:e1006899.
Rio RVM, Jozwick AKS, Savage AF, Sabet A, Vigneron A, Wu Y, et al. Mutualist-Provisioned Resources Impact Vector Competency. mBio. 2019;10:e00018-19.
Michalkova V, Benoit JB, Weiss BL, Attardo GM, Aksoy S. Vitamin B6 generated by obligate symbionts is critical for maintaining proline homeostasis and fecundity in tsetse flies. Appl Environ Microbiol [Internet]. 2014;80:5844–53. Available from: http://dx.doi.org/10.1128/AEM.01150-14
Michalkova V, Benoit JB, Attardo GM, Medlock J, Aksoy S. Amelioration of reproduction-associated oxidative stress in a viviparous insect is critical to prevent reproductive senescence. PLoS One [Internet]. 2014;9:e87554-. Available from: DOI:10.1371/journal.pone.0087554
Moloo SK, Kabata JM, Waweru F, Gooding RH. Selection of susceptible and refractory lines of Glossina morsitans centralis for Trypanosoma congolense infection and their susceptibility to different pathogenic Trypanosoma species. Med Vet Entomol. 1998;12:391–8.
Farikou O, Thevenon S, Njiokou F, Allal F, Cuny G, Geiger A. Genetic diversity and population structure of the secondary symbiont of tsetse flies, Sodalis glossinidius, in sleeping sickness foci in Camerooon. PLoS Negl Trop Dis [Internet]. 2011;5:e1281-. Available from: doi:10.1371/journal.pntd.0001281
Soumana IH, Simo G, Njiokou F, Tchicaya B, Abd-Alla AMM, Cuny G, et al. The bacterial flora of tsetse fly midgut and its effect on trypanosome transmission. J Invertebr Pathol [Internet]. 2013;112:S89–93. Available from: http://dx.doi.org/10.1016/j.jip.2012.03.029
Aksoy E, Telleria E, Echodu R, Wu Y, Okedi LM, Weiss BL, et al. Analysis of multiple tsetse fly populations in Uganda reveals limited diversity and species-specific gut microbiota. Appl Environ Microbiol [Internet]. 2014;80:4301–12. Available from: DOI:10.1128/AEM.00079-14
Griffith BC, Weiss BL, Aksoy E, Mireji PO, Auma JE, Wamwiri FN, et al. Analysis of the gut-specific microbiome from field-captured tsetse flies, and its potential relevance to host trypanosome vector competence. BMC Microbiol [Internet]. 2018 [cited 2020 Nov 14];18:146. Available from: https://doi.org/10.1186/s12866-018-1284-7
Weiss BL, Maltz MA, Vigneron A, Wu Y, Walter KS, O’Neill MB, et al. Colonization of the tsetse fly midgut with commensal Kosakonia cowanii Zambiae inhibits trypanosome infection establishment. PLoS Pathog. 2019;15:e1007470.
Leak SGA. Tsetse biology and ecology: their role in the epidemiology and control of trypanosomosis. Wallingford: CABI Publishing; 1998.
Simarro PP, Louis FJ, Jannin J. Sleeping sickness, forgotten illness: what are the impact in the field? MedTrop. 2003;63:231–5.
Dyck VA, Hendrichs J, Robinson AS. Sterile Insect Technique. Principles and Practice in Area-Wide Integrated Pest Management. Second Edition. CRC Press, Boco Raton, FL; 2021.
Feldmann U, Dyck VA, Mattioli R, Jannin J, Vreysen MJB. Impact of tsetse fly eradication programmes using the sterile insect technique. In: Dyck VA, Hendrichs JP, Robinson AS, editors. Sterile Insect Tech Princ Pract Area-Wide Integr Pest Manag [Internet]. 2nd ed. Boca Raton, FL: CRC Press; 2021. p. 701–30. Available from: https://www.taylorfrancis.com/chapters/impact-tsetse-fly-eradication-programmes-using-sterile-insect-technique-feldmann-dyck-mattioli-jannin-vreysen/e/10.1201/9781003035572-32
Vreysen MJB, Saleh KM, Ali MY, Abdulla AM, Zhu Z-R, Juma KG, et al. Glossina austeni (Diptera: Glossinidae) eradicated on the island of Unguja, Zanzibar, using the sterile insect technique. J Econ Entomol [Internet]. 2000;93:123–35. Available from: DOI:10.1603/0022-0493-93.1.123
Hendrichs MA, Wornoayporn V, Katsoyannos BI, Hendrichs JP. Quality control method to measure predator evasion in wild and mass-reared Mediterranean fruit flies (Diptera: Tephritidae). Fla Entomol. 2007;90:64–70.
Vreysen MJB, Abd-Alla AMM, Bourtzis K, Bouyer J, Caceres C, de Beer C, et al. The Insect Pest Control Laboratory of the Joint FAO/IAEA Programme: Ten Years (2010-2020) of Research and Development, Achievements and Challenges in Support of the Sterile Insect Technique. Insects. 2021;12:346.
Attardo GM, Lohs C, Heddi A, Alam UH, Yildirim S, Aksoy S. Analysis of milk gland structure and function in Glossina morsitans: Milk protein production, symbiont populations and fecundity. J Insect Physiol [Internet]. 2008;54:1236–42. Available from: http://www.sciencedirect.com/science/journal/00221910
Belda E, Moya A, Bentley S, Silva FJ. Mobile genetic element proliferation and gene inactivation impact over the genome structure and metabolic capabilities of Sodalis glossinidius, the secondary endosymbiont of tsetse flies. BMC Genomics. 2010;11:449-.
International Glossina Genome Initiative. Genome sequence of the tsetse fly (Glossina morsitans): vector of African trypanosomiasis. Science [Internet]. 2014;344:380–6. Available from: DOI:10.1126/science.1249656
Abd-Alla AMM, Kariithi HM, Parker AG, Robinson AS, Kiflom M, Bergoin M, et al. Dynamics of the salivary gland hypertrophy virus in laboratory colonies of Glossina pallidipes (Diptera: Glossinidae). Virus Res [Internet]. 2010;150:103–10. Available from: http://dx.doi.org/10.1016/j.virusres.2010.03.001
Aksoy S. Tsetse- A haven for microorganisms. Parasitol Today. 2000;16:114–8.
Aksoy S, Weiss B, Attardo GM. Paratransgenesis applied for control of tsetse transmitted sleeping sickness. In: Serap Aksoy, editor. 627th ed. New York: Springer Science-Business Media, LLC Landes Bioscience (www.spinger.com); 2008. p. 35–48.
Alam U, Medlok J, Brelsfoard C, Pais R, Lohs C, Balmand S, et al. Wolbachia symbiont infections induce strong cytoplasmic incompatibility in the tsetse fly Glossina morsitans. PLoS Pathog [Internet]. 2011;7:e1002415-. Available from: doi:10.1371/journal.ppat.1002415
Doudoumis V, Alam U, Aksoy E, Abd-Alla AMM, Tsiamis G, Brelsfoard C, et al. Tsetse-Wolbachia symbiosis: Comes of age and has great potential for pest and disease control. J Invertebr Pathol [Internet]. 2013;112:S94–103. Available from: http://dx.doi.org/10.1016/j.jip.2012.05.010
Lietze VU, Abd-Alla AMM, Vreysen MJB, Geden CJ, Boucias DG. Salivary gland hypertrophy viruses: a novel group of insect pathogenic viruses. Annu Rev Entomol [Internet]. 2010;56:63–80. Available from: doi:10.1146/annurev-ento-120709-144841
Kariithi HM, Ahmadi M, Parker AG, Franz G, Ros VID, Haq I, et al. Prevalence and genetic variation of salivary gland hypertrophy virus in wild populations of the tsetse fly Glossina pallidipes from southern and eastern Africa. J Invertebr Pathol [Internet]. 2013;112:S123–32. Available from: DOI: 10.1016/j.jip.2012.04.016
Abd-Alla AMM, Kariithi HM, Bergoin M. Managing Pathogens in Insect Mass-Rearing for the Sterile Insect Technique, with the Tsetse Fly Salivary Gland Hypertrophy Virus as an Example. In: Dyck VA, Hendrichs JP, Robinson AS, editors. Sterile Insect Tech Princ Pract Area-Wide Integr Pest Manag [Internet]. 2nd ed. Boca Raton, FL: CRC Press; 2021. p. 317–54. Available from: https://www.taylorfrancis.com/chapters/managing-pathogens-insect-mass-rearing-sterile-insect-technique-tsetse-fly-salivary-gland-hypertrophy-virus-example-abd-alla-kariithi-bergoin/e/10.1201/9781003035572-10
Demirbas-Uzel G, Augustinos AA, Doudoumis V, Parker AG, Tsiamis G, Bourtzis K, et al. Interactions between tsetse endosymbionts and Glossina pallidipes salivary gland hypertrophy virus in Glossina hosts. Front Microbiol [Internet]. 2021;12:1295. Available from: https://www.frontiersin.org/article/10.3389/fmicb.2021.653880
Baker RD, Maudlin I, Milligan PJM, Molyneux DH, Welburn SC. The possible role of Rickettsia-like organisms in trypanosomiasis epidemiology. Parasitology. 1990;100:209–17.
Soumana IH, Berthier D, Tchicaya B, Thevenon S, Njiokou F, Cuny G, et al. Population dynamics of Glossina palpalis gambiensis symbionts, Sodalis glossinidius, and Wigglesworthia glossinidia, throughout host-fly development. Infect Genet Evol [Internet]. 2013;13:41–8. Available from: DOI:10.1016/j.meegid.2012.10.003
Dennis JW, Durkin SM, Downie JEH, Hamill LC, Anderson NE, MacLeod ET. Sodalis glossinidius prevalence and trypanosome presence in tsetse from Luambe National Park, Zambia. 2014;11.
Meki IK, Kariithi HM, Ahmadi M, Parker AG, Vreysen MJB, Vlak JM, et al. Hytrosavirus genetic diversity and eco-regional spread in Glossina species. BMC Microbiol [Internet]. 2018 [cited 2019 May 29];18:143. Available from: https://doi.org/10.1186/s12866-018-1297-2
Ouedraogo GMS, Demirbas-Uzel G, Rayaisse J-B, Gimonneau G, Traore AC, Avgoustinos A, et al. Prevalence of trypanosomes, salivary gland hypertrophy virus and Wolbachia in wild populations of tsetse flies from West Africa. BMC Microbiol. 2018;18:153.
Abd-Alla AMM, Cousserans F, Parker A, Bergoin M, Chiraz J, Robinson A. Quantitative PCR analysis of the salivary gland hypertrophy virus (GpSGHV) in a laboratory colony of Glossina pallidipes. Virus Res [Internet]. 2009;139:48–53. Available from: http://dx.doi.org/10.1016/j.virusres.2008.10.006
Demirbas-Uzel G, De Vooght L, Parker AG, Vreysen MJB, Mach RL, Van Den Abbeele J, et al. Combining paratransgenesis with SIT: impact of ionizing radiation on the DNA copy number of Sodalis glossinidius in tsetse flies. BMC Microbiol [Internet]. 2018 [cited 2019 May 29];18:160. Available from: https://doi.org/10.1186/s12866-018-1283-8
R Core Team. R: A Language and Environment for Statistical Computing [Internet]. Vienna, Austria: R Foundation for Statistical Computing; 2021. Available from: https://www.R-project.org/
Baier T, Neuwirth E. Excel:: COM :: R. Comput Stat [Internet]. 2007;22:91–108. Available from: DOI 10.1007/s00180-007-0023-6
RStudio Team. RStudio: Integrated Development Environment for R [Internet]. Boston, MA: RStudio, Inc.; 2016. Available from: http://www.rstudio.com/
Wickham H. ggplot2: Elegant Graphics for Data Analysis [Internet]. Springer-Verlag New York; 2016. Available from: https://ggplot2.tidyverse.org
Sarkar D. Lattice: multivariate data visualization with R. Springer Science & Business Media; 2008.
Fox J, Weisberg S. An R Companion to Applied Regression, Second Edition [Internet]. Third. Thousand Oaks, CA: Sage; 2019. Available from: https://socialsciences.mcmaster.ca/jfox/Books/Companion/
Jeffrey B. Arnold. ggthemes: Extra Themes, Scales and Geoms for “ggplot2”. R package version 4.2.4. [Internet]. 2021. Available from: https://CRAN.R-project.org/package=ggthemes
Venables WN, Ripley BD. Modern Applied Statistics with S [Internet]. Fourth. New York: Springer; 2002. Available from: http://www.stats.ox.ac.uk/pub/MASS4/
Clarke KR, Gorley RN. Getting started with PRIMER v7 [Internet]. 2016. Available from: http://updates.primer-e.com/primer7/manuals/Getting_started_with_PRIMER_7.pdf
Anderson MJ. A new method for non-parametric multivariate analysis of variance. Austral Ecol [Internet]. 2001 [cited 2020 Nov 23];26:32–46. Available from: https://onlinelibrary.wiley.com/doi/abs/10.1111/j.1442-9993.2001.01070.pp.x
Alam U, Hyseni C, Symula RE, Brelsfoard C, Wu Y, Kruglov O, et al. Implications of microfauna-host interactions for trypanosome transmission dynamics in Glossina fuscipes fuscipes in Uganda. Appl Env Microbiol. 2012;78:4627–37.
Hedges LM, Brownlie JC, O’Neill SL, Johnson KN. Wolbachia and virus protection in insects. Science [Internet]. 2008;322:702-. Available from: DOI: 10.1126/science.1162418
Teixeira L, Ferreira A, Ashburner M. The bacterial symbiontWolbachiainduces resistance to RNA viral infections inDrosophila melanogaster. PLoS Biol. 2008;6:e1000002-.
Johnson KN. The Impact of Wolbachia on Virus Infection in Mosquitoes. Viruses [Internet]. Multidisciplinary Digital Publishing Institute; 2015 [cited 2020 Aug 2];7:5705–17. Available from: https://www.mdpi.com/1999-4915/7/11/2903
Graham RI, Grzywacz D, Mushobozi WL, Wilson K. Wolbachia in a major African crop pest increases susceptibility to viral disease rather than protects. Ecol Lett [Internet]. 2012;15:993–1000. Available from: DOI:10.1111/j.1461-0248.2012.01820.x
Dodson BL, Hughes GL, Paul O, Matacchiero AC, Kramer LD, Rasgon JL. Wolbachia enhances West Nile virus (WNV) infection in the mosquito Culex tarsalis. PLoS Negl Trop Dis [Internet]. 2014;8:e2965-. Available from: DOI:10.1371/journal.pntd.0002965
Lu P, Bian G, Pan X, Xi Z. Wolbachia Induces Density-Dependent Inhibition to Dengue Virus in Mosquito Cells. PLoS Negl Trop Dis [Internet]. Public Library of Science; 2012 [cited 2020 Aug 2];6:e1754. Available from: https://journals.plos.org/plosntds/article?id=10.1371/journal.pntd.0001754
Parry R, Bishop C, De Hayr L, Asgari S. Density-dependent enhanced replication of a densovirus in Wolbachia-infected Aedes cells is associated with production of piRNAs and higher virus-derived siRNAs. Virology [Internet]. 2019 [cited 2020 Aug 2];528:89–100. Available from: http://www.sciencedirect.com/science/article/pii/S0042682218303702
Weiss BL, Wang J, Aksoy S. Tsetse immune system maturation requires the presence of obligate symbionts in larvae. PLoS Biol. 2011;9:e1000619-.

AdditionalTableS1.docx
AdditionalTableS2.xlsx
AdditionalfileS1.docx
FigureS1.tif
Supplementary Figure 1. Prevalence of GpSGHV and Wolbachia co-infection in natural tsetse populations. A: in all tsetse species, B: in each tsetse species. GpSGHV and Wolbachia prevalence was determined by PCR as described previously [7,47].
FigureS2.tif
Supplementary Figure 2. Density levels of GpSGHV (A), Wolbachia (B), Wigglesworthia (C) and Sodalis (D) in tsetse flies with different GpSGHV and Wolbachia infection status. The copy number was determined by qPCR. Values indicated by a different small letter differ significantly at the 5% level. W+/V+ : flies infected with both Wolbachia and GpSGHV; W+/V- : flies infected only with Wolbachia, and W-/V-+ : flies infected only with GpSGHV.
FigureS3.tif
Supplementary Figure 3. Density levels of GpSGHV (A), Wolbachia (B), Wigglesworthia (C) and Sodalis (D) in tsetse flies collected from different countries. The copy number was determined by qPCR. Values indicated by a different small letter differ significantly at the 5% level.
FigureS4.tif
Supplementary Figure 4. Impact of GpSGHV and Wolbachia co-infection (W+/V+) on the density levels of GpSGHV (A), Wolbachia (B), Wigglesworthia (C) and Sodalis (D) in different tsetse species. The copy number was determined by qPCR. Values indicated by the same lower-case letter do not differ significantly at the 5% level.
FigureS5.tif
Supplementary Figure 5. Relative density of GpSGHV, Wigglesworthia, Sodalis and Wolbachia in G. austeni and G. m. morsitans field collected tsetse flies. The density of GpSGHV and tsetse symbionts were analyzed by qPCR. Data were transformed to square root and averaged based on countries and species (A), countries and infection status (Sample) (B), and countries, species, and infection status (Sample) (C). The top and the left of the graph indicate the group averaged Bray-Curtis similarity.
Graphicalabstract.tif

Download PDF

Version 1

posted

You are reading this older preprint version

Read the latest preprint version →

Interactions Between Glossina Pallidipes Salivary Gland Hypertrophy Virus and Tsetse Endosymbionts in Wild Tsetse Populations

Archived Versions:

Version 1

Abstract

Figures

Introduction

Material And Methods

Statistical analysis

Results

Prevalence of coinfection with GpSGHV and Wolbachia in wild tsetse flies

Impact of co-infection (W⁺/V⁺) on GpSGHV, Wolbachia, Sodalis and Wigglesworthia infection density

GpSGHV density

Wolbachia density

Interaction between GpSGHV infection and Wolbachia, Wigglesworthia, and Sodalis infection

Discussion

Conclusions

Declarations

References

Supplementary Files

Archived Versions:

Version 1

Interactions Between Glossina Pallidipes Salivary Gland Hypertrophy Virus and Tsetse Endosymbionts in Wild Tsetse Populations

Archived Versions:

Version 1

Abstract

Figures

Introduction

Material And Methods

Statistical analysis

Results

Prevalence of coinfection with GpSGHV and Wolbachia in wild tsetse flies

Impact of co-infection (W+/V+) on GpSGHV, Wolbachia, Sodalis and Wigglesworthia infection density

GpSGHV density

Wolbachia density

Interaction between GpSGHV infection and Wolbachia, Wigglesworthia, and Sodalis infection

Discussion

Conclusions

Declarations

References

Supplementary Files

Archived Versions:

Version 1

Impact of co-infection (W⁺/V⁺) on GpSGHV, Wolbachia, Sodalis and Wigglesworthia infection density