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    Box plots of number of Plasmodium yoelii oocysts per mosquito in oocyst-positive Anopheles stephensi fed on Echinostoma caproniP. yoelii co-infected mice (E+M) (n = 193 mosquitoes) or P. yoelii only–infected mice (M) (n = 167 mosquitoes) five days post-malaria infection (aggregate of four experiments). Boxes, with medians indicated, contain the middle 50% of observations (the interquartile range [IQR]), whiskers indicate the smallest and largest observations within the lower and upper fence (± 1.5 × IQR), and outliers are observations outside the upper fence. Geometric mean number of oocysts were 19.2 (E+M) and 10.5 (M) (P = 0.004, by Student’s t-test of log-transformed values.)

  • 1

    Karunaweera ND, Carter R, Grau GE, Kwiatkowski D, del Giudice G, Mendis KN, 1992. Tumour necrosis factor-dependent parasite-killing effects during paroxysms in non-immune Plasmodium vivax malaria patients. Clin Exp Immunol 88 :499–505.

    • Search Google Scholar
    • Export Citation
  • 2

    Naotunne TS, Karunaweera ND, Del Giudice G, Kularatne MU, Grau GE, Carter R, Mendis KN, 1991. Cytokines kill malaria parasites during infection crisis: extracellular complementary factors are essential. J Exp Med 173 :523–529.

    • Search Google Scholar
    • Export Citation
  • 3

    Kaushal DC, Carter R, Rener J, Grotendorst CA, Miller LH, Howard RJ, 1983. Monoclonal antibodies against surface determinants on gametes of Plasmodium gallinaceum block transmission of malaria parasites to mosquitoes. J Immunol 131 :2557–2562.

    • Search Google Scholar
    • Export Citation
  • 4

    Rener J, Graves PM, Carter R, Williams JL, Burkot TR, 1983. Target antigens of transmission-blocking immunity on gametes of Plasmodium falciparum.J Exp Med 158 :976–981.

    • Search Google Scholar
    • Export Citation
  • 5

    Naotunne TD, Rathnayake KD, Jayasinghe A, Carter R, Mendis KN, 1990. Plasmodium cynomolgi: serum-mediated blocking and enhancement of infectivity to mosquitoes during infections in the natural host, Macaca sinica.Exp Parasitol 71 :305–313.

    • Search Google Scholar
    • Export Citation
  • 6

    Peiris JS, Premawansa S, Ranawaka MB, Udagama PV, Munasinghe YD, Nanayakkara MV, Gamage CP, Carter R, David PH, Mendis KN, 1988. Monoclonal and polyclonal antibodies both block and enhance transmission of human Plasmodium vivax malaria. Am J Trop Med Hyg 39 :26–32.

    • Search Google Scholar
    • Export Citation
  • 7

    Butcher GA, Sinden RE, Billker O, 1996. Plasmodium berghei: infectivity of mice to Anopheles stephensi mosquitoes. Exp Parasitol 84 :371–379.

    • Search Google Scholar
    • Export Citation
  • 8

    de Silva NR, Brooker S, Hotez PJ, Montresor A, Engels D, Savioli L, 2003. Soil-transmitted helminth infections: updating the global picture. Trends Parasitol 19 :547–551.

    • Search Google Scholar
    • Export Citation
  • 9

    Mwangi TW, Bethony JM, Brooker S, 2006. Malaria and helminth interactions in humans: an epidemiological viewpoint. Ann Trop Med Parasitol 100 :551–570.

    • Search Google Scholar
    • Export Citation
  • 10

    Buck AA, Anderson RI, MacRae AA, 1978. Epidemiology of poly-parasitism. IV. combined effects on the state of health. Trop Med Parasitol 29 :253–268.

    • Search Google Scholar
    • Export Citation
  • 11

    Nacher M, Singhasivanon P, Yimsamran S, Manibunyong W, Thanyavanich N, Wuthisen R, Looareesuwan S, 2002. Intestinal helminth infections are associated with increased incidence of Plasmodium falciparum malaria in Thailand. J Parasitol 88 :55–58.

    • Search Google Scholar
    • Export Citation
  • 12

    Spiegel A, Tall A, Raphenon G, Trape JF, Druilhe P, 2003. Increased frequency of malaria attacks in subjects co-infected by intestinal worms and Plasmodium falciparum malaria. Trans R Soc Trop Med Hyg 97 :198–199.

    • Search Google Scholar
    • Export Citation
  • 13

    Le Hesran JY, Akiana J, Ndiaye EH, Dia M, Senghor P, Konate L, 2004. Severe malaria attack is associated with high prevalence of Ascaris lumbricoides infection among children in rural Senegal. Trans R Soc Trop Med Hyg 98 :397–399.

    • Search Google Scholar
    • Export Citation
  • 14

    Sokhna C, Le Hesran JY, Mbaye PA, Akiana J, Camara P, Diop M, Ly A, Druilhe P, 2004. Increase of malaria attacks among children presenting concomitant infection by Schistosoma mansoni in Senegal. Malar J 3 :43.

    • Search Google Scholar
    • Export Citation
  • 15

    Nacher M, Gay F, Singhasivanon P, Krudsood S, Treeprasertsuk S, Mazier D, Vouldoukis I, Looareesuwan S, 2000. Ascaris lumbricoides infection is associated with protection from cerebral malaria. Parasite Immunol 22 :107–113.

    • Search Google Scholar
    • Export Citation
  • 16

    Nacher M, Singhasivanon P, Silachamroon U, Treeprasertsuk S, Vannaphan S, Traore B, Gay F, Looareesuwan S, 2001. Helminth infections are associated with protection from malaria-related acute renal failure and jaundice in Thailand. Am J Trop Med Hyg 65 :834–836.

    • Search Google Scholar
    • Export Citation
  • 17

    Lyke KE, Dicko A, Dabo A, Sangare L, Kone A, Coulibaly D, Guindo A, Traore K, Daou M, Diarra I, Sztein MB, Plowe CV, Doumbo OK, 2005. Association of Schistosoma haematobium infection with protection against acute Plasmodium falciparum malaria in Malian children. Am J Trop Med Hyg 73 :1124–1130.

    • Search Google Scholar
    • Export Citation
  • 18

    Briand V, Watier L, Hesran JY, Garcia A, Cot M, 2005. Coin-fection with Plasmodium falciparum and Schistosoma haematobium: protective effect of schistosomiasis on malaria in Senegalese children? Am J Trop Med Hyg 72 :702–707.

    • Search Google Scholar
    • Export Citation
  • 19

    Brutus L, Watier L, Briand V, Hanitrasoamampionona V, Razanatsoarilala H, Cot M, 2006. Parasitic co-infections: Does As-caris lumbricoides protect against Plasmodium falciparum infection? Am J Trop Med Hyg 75 :194–198.

    • Search Google Scholar
    • Export Citation
  • 20

    Lwin M, Last C, Targett GA, Doenhoff MJ, 1982. Infection of mice concurrently with Schistosoma mansoni and rodent malarias: contrasting effects of patent S. mansoni infections on Plasmodium chabaudi, P. yoelii and P. berghei.Ann Trop Med Parasitol 76 :265–273.

    • Search Google Scholar
    • Export Citation
  • 21

    Christensen NØ, Furu P, Kurtzhals J, Odaibo A, 1988. Heterologous synergistic interactions in concurrent experimental infection in the mouse with Schistosoma mansoni, Echinostoma revolutum, Plasmodium yoelii, Babesia microti, and Trypanosoma brucei.Parasitol Res 74 :544–551.

    • Search Google Scholar
    • Export Citation
  • 22

    Noland GS, Graczyk TK, Fried B, Fitzgerald EJ, Kumar N, 2005. Exacerbation of Plasmodium yoelii malaria in Echinostoma caproni infected mice and abatement through anthelmintic treatment. J Parasitol 91 :944–948.

    • Search Google Scholar
    • Export Citation
  • 23

    Helmby H, Kullberg M, Troye-Blomberg M, 1998. Altered immune responses in mice with concomitant Schistosoma mansoni and Plasmodium chabaudi infections. Infect Immun 66 :5167–5174.

    • Search Google Scholar
    • Export Citation
  • 24

    Yoshida A, Maruyama H, Kumagai T, Amano T, Kobayashi F, Zhang M, Himeno K, Ohta N, 2000. Schistosoma mansoni infection cancels the susceptibility to Plasmodium chabaudi through induction of type 1 immune responses in A/J mice. Int Immunol 12 :1117–1125.

    • Search Google Scholar
    • Export Citation
  • 25

    Su Z, Segura M, Morgan K, Loredo-Osti JC, Stevenson MM, 2005. Impairment of protective immunity to blood-stage malaria by concurrent nematode infection. Infect Immun 73 :3531–3539.

    • Search Google Scholar
    • Export Citation
  • 26

    Yan Y, Inuo G, Akao N, Tsukidate S, Fujita K, 1997. Down-regulation of murine susceptibility to cerebral malaria by inoculation with third-stage larvae of the filarial nematode Brugia pahangi.Parasitology 114 :333–338.

    • Search Google Scholar
    • Export Citation
  • 27

    Graham AL, Lamb TJ, Read AF, Allen JE, 2005. Malaria-filaria coinfection in mice makes malarial disease more severe unless filarial infection achieves patency. J Infect Dis 191 :410–421.

    • Search Google Scholar
    • Export Citation
  • 28

    Bastien P, Landau I, Baccam D, 1987. Inhibition of the infectivity of Plasmodium gametocytes by the serum of the parasite host. Perfecting an experimental model. Ann Parasitol Hum Comp 62 :195–208.

    • Search Google Scholar
    • Export Citation
  • 29

    Nacher M, Singhasivanon P, Silachamroon U, Treeprasertsu S, Krudsood S, Gay F, Mazier D, Looareesuwan S, 2001. Association of helminth infections with increased gametocyte carriage during mild falciparum malaria in Thailand. Am J Trop Med Hyg 65 :644–647.

    • Search Google Scholar
    • Export Citation
  • 30

    Mellor PS, Boorman J, 1980. Multiplication of bluetongue virus in Culicoides nubeculosus (Meigen) simultaneously infected with the virus and the microfilariae of Onchocerca cervicalis (Railliet & Henry). Ann Trop Med Parasitol 74 :463–469.

    • Search Google Scholar
    • Export Citation
  • 31

    Paulson SL, Poirier SJ, Grimstad PR, Craig GB Jr, 1992. Vector competence of Aedes hendersoni (Diptera: Culicidae) for la crosse virus: Lack of impaired function in virus-infected salivary glands and enhanced virus transmission by sporozoite-infected mosquitoes. J Med Entomol 29 :483–488.

    • Search Google Scholar
    • Export Citation
  • 32

    Turell MJ, Mather TN, Spielman A, Bailey CL, 1987. Increased dissemination of dengue 2 virus in Aedes aegypti associated with concurrent ingestion of microfilariae of Brugia malayi.Am J Trop Med Hyg 37 :197–201.

    • Search Google Scholar
    • Export Citation
  • 33

    Vaughan JA, Turell MJ, 1996. Dual host infections: enhanced infectivity of eastern equine encephalitis virus to Aedes mosquitoes mediated by Brugia microfilariae. Am J Trop Med Hyg 54 :105–109.

    • Search Google Scholar
    • Export Citation
  • 34

    Vaughan JA, Turell MJ, 1996. Facilitation of rift valley fever virus transmission by Plasmodium berghei sporozoites in Anopheles stephensi mosquitoes. Am J Trop Med Hyg 55 :407–409.

    • Search Google Scholar
    • Export Citation
  • 35

    Vaughan JA, Trpis M, Turell MJ, 1999. Brugia malayi microfilariae (Nematoda: Filaridae) enhance the infectivity of Venezuelan equine encephalitis virus to Aedes mosquitoes (Diptera: Culicidae). J Med Entomol 36 :758–763.

    • Search Google Scholar
    • Export Citation
  • 36

    Zytoon EM, el Belbasi HI, Matsumura T, 1993. Mechanism of increased dissemination of Chikungunya virus in Aedes albopictus mosquitoes concurrently ingesting microfilariae of Dirofilaria immitis.Am J Trop Med Hyg 49 :201–207.

    • Search Google Scholar
    • Export Citation
  • 37

    Burkot TR, Molineaux L, Graves PM, Paru R, Battistutta D, Dagoro H, Barnes A, Wirtz RA, Garner P, 1990. The prevalence of naturally acquired multiple infections of Wuchereria bancrofti and human malarias in anophelines. Parasitology 100 :369–375.

    • Search Google Scholar
    • Export Citation
  • 38

    Munderloh UG, Kurtti TJ, 1987. The infectivity and purification of cultured Plasmodium berghei ookinetes. J Parasitol 73 :919–923.

  • 39

    Ponnudurai T, Lensen AH, Van Gemert GJ, Bensink MP, Bolmer M, Meuwissen JH, 1989. Infectivity of cultured Plasmodium falciparum gametocytes to mosquitoes. Parasitology 98 :165–173.

    • Search Google Scholar
    • Export Citation
  • 40

    Haji H, Smith T, Charlwood JD, Meuwissen JH, 1996. Absence of relationships between selected human factors and natural infectivity of Plasmodium falciparum to mosquitoes in an area of high transmission. Parasitology 113 :425–431.

    • Search Google Scholar
    • Export Citation
  • 41

    Sinden RE, Butcher GA, Billker O, Fleck SL, 1996. Regulation of infectivity of Plasmodium to the mosquito vector. Adv Parasitol 38 :53–117.

    • Search Google Scholar
    • Export Citation
  • 42

    Gautret P, Gantier JC, Baccam D, Miltgen F, Saulai M, Chabaud AG, Landau I, 1996. The gametocytes of Plasmodium vinckei petteri, their morphological stages, periodicity and infectivity. Int J Parasitol 26 :1095–1101.

    • Search Google Scholar
    • Export Citation
  • 43

    Gautret P, Miltgen F, Chabaud AG, Landau I, 1996. Synchronized Plasmodium yoelii yoelii: pattern of gametocyte production, sequestration and infectivity. Parassitologia 38 :575–577.

    • Search Google Scholar
    • Export Citation
  • 44

    Gautret P, Miltgen F, Gantier JC, Chabaud AG, Landau I, 1996. Enhanced gametocyte formation by Plasmodium chabaudi in immature erythrocytes: pattern of production, sequestration, and infectivity to mosquitoes. J Parasitol 82 :900–906.

    • Search Google Scholar
    • Export Citation
  • 45

    Wargo AR, Randle N, Chan BH, Thompson J, Read AF, Babiker HA, 2006. Plasmodium chabaudi: reverse transcription PCR for the detection and quantification of transmission stage malaria parasites. Exp Parasitol 112 :13–20.

    • Search Google Scholar
    • Export Citation
  • 46

    Hotez PJ, Brooker S, Bethony JM, Bottazzi ME, Loukas A, Xiao S, 2004. Hookworm infection. N Engl J Med 351 :799–807.

  • 47

    Friedman JF, Kanzaria HK, McGarvey ST, 2005. Human schistosomiasis and anemia: The relationship and potential mechanisms. Trends Parasitol 21 :386–392.

    • Search Google Scholar
    • Export Citation
  • 48

    Price R, Nosten F, Simpson JA, Luxemburger C, Phaipun L, ter Kuile F, van Vugt M, Chongsuphajaisiddhi T, White NJ, 1999. Risk factors for gametocyte carriage in uncomplicated falciparum malaria. Am J Trop Med Hyg 60 :1019–1023.

    • Search Google Scholar
    • Export Citation
  • 49

    Wisnewski N, Fried B, Halton DW, 1986. Growth and feeding of Echinostoma revolutum on the chick chorioallantois and in the domestic chick. J Parasitol 72 :684–689.

    • Search Google Scholar
    • Export Citation
  • 50

    Taylor PJ, Hurd H, 2001. The influence of host haematocrit on the blood feeding success of Anopheles stephensi: implications for enhanced malaria transmission. Parasitology 122 :491–496.

    • Search Google Scholar
    • Export Citation
  • 51

    Murray J, Murray A, Murray M, Murray C, 1978. The biological suppression of malaria: an ecological and nutritional interrelationship of a host and two parasites. Am J Clin Nutr 31 :1363–1366.

    • Search Google Scholar
    • Export Citation
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ENHANCED MALARIA PARASITE TRANSMISSION FROM HELMINTH CO-INFECTED MICE

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  • 1 Department of Molecular Microbiology and Immunology, Johns Hopkins Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland; Department of Biology, Lafayette College, Easton, Pennsylvania

Helminth infections are prevalent in malaria-endemic areas, yet the potential for helminths to alter malaria transmission has not been closely examined. We used the Echinostoma caproniPlasmodium yoelii murine model of co-infection to assess the impact of helminth co-infection on malaria transmission. In four replicate experiments, Anopheles stephensi mosquitoes exposed to co-infected mice five days post-malaria infection had a higher rate of infectivity (80.1%, n = 241) than those exposed to malaria only–infected mice (72.0%, n = 232, P = 0.039). Intensity of malaria parasite transmission was also greater, with approximately two-fold more oocysts (geometric mean = 19.2 versus 10.5, P = 0.004) and an increase in sporozoite burden observed in mosquitoes exposed to co-infected mice. Malaria parasite prevalence and anemia were similar between co-infected and malaria only–infected mice, which suggested that enhanced malaria parasite transmission was due to helminth-induced modulation of host responses.

INTRODUCTION

Malaria, a disease that affects more than 300 million people annually, is transmitted between vertebrate hosts by female Anopheles mosquitoes. Infectivity of the transmissible form of the parasite, the gametocyte, can be influenced by various vertebrate host factors, including cytokines, antibodies to parasites, and non-specific alterations to host physiology. Th1 cytokines, specifically tumor necrosis factor-α and interferon- γ, have been shown to reduce transmission of Plasmodium vivax by neutralizing infectivity of gametocytes in conjunction with other serum factors produced during paroxysm.1,2 Antibodies directed against sexual stage parasites can likewise reduce transmission,3,4 however, paradoxically, some sexual stage-specific antibodies can also enhance malaria transmission.5,6 Furthermore, in rodent malaria infections, decreases in host pH and bicarbonate levels have been shown to inhibit transmission independently of other serum derived factors by preventing exflagellation of microgametes.7

Approximately two billion people in malaria-endemic areas also have various helminth infections.8 Concurrent infection with malaria and helminths is common,9 and numerous studies have documented significant interactions, both synergistic1014 and protective,1519 in human populations. In animal models, similarly mixed profiles have been described. Patent helminth infections tend to exacerbate normally non-lethal P. yoelii2022 and P. chabaudi2325 infections, and provide protection against some, but not all, lethal P. yoelii, P. chabaudi, and P. berghei infections.20,22,24,26 Although the mechanisms contributing to such outcomes are not clearly defined, significant modulation of Th1 cytokines and anti-malaria antibodies has been reported during co-infection.2325,27 We therefore hypothesized that such alterations to host response during concomitant helminth infection may also modulate the intensity of malaria parasite transmission. Additionally, because helminth infection has been found to increase the duration of malaria parasitemia,20,22 we sought to determine whether chronic helminth infection extended the permissive window of P. yoelii transmission, which is normally limited to the first five days of infection.28

One previous study has suggested that helminths may modulate malaria transmission potential because helminth-infected individuals with mild P. falciparum malaria were more likely to carry gametocytes than patients with malaria alone.29 However, the effect on actual transmission to mosquitoes has never been addressed. We used an established rodent model of co-infection,22 using an intestinal trematode, Echinostoma caproni, and a non lethal rodent malaria parasite, Plasmodium yoelii, to evaluate the impact of helminth infection on malaria transmission by comparing malaria parasite burdens in mosquitoes exposed to helminth co-infected or malaria only–infected mice.

MATERIALS AND METHODS

Mice and parasites.

Four- to six-week old male BALB/c mice were obtained from the National Cancer Institute (Bethesda, MD) and maintained in a pathogen-free micro-isolation facility in accordance with the National Institutes of Health guidelines for the humane use of laboratory animals. Mice were infected with 10–15 E. caproni metacercarial cysts, as described previously,22 and E. caproni infections were monitored by microscopic examination of weekly fecal collections. Approximately four weeks after helminth infection, groups of 3–5 E. caproni-infected mice and age- and sex-matched worm-free controls were infected with 1 × 105 P. yoelii parasites (17X non-lethal strain) by intraperitoneal injection of infected erythrocytes. Each of four independent replicate experiments was initiated from the same frozen parasite pool and passaged through one donor mouse prior to experimental infections. Parasitemia and gametocytemia were determined by counting 1 × 103 and 1 × 104 erythrocytes, respectively, in Giemsa-stained thin films of mouse tail blood. Anemia was determined by measuring packed cell volume of blood drawn by tail bleed into micro-capillary tubes.

Mosquito infections.

At days five, eight, and thirteen post-malaria infection, mice were anesthetized with an intraperitoneally injection of katamine (Ketaject, 100 mg/kg; Phoenix Pharmaceuticals, St. Joseph, MO) and acepromazine (5 mg/ kg; Henry Schein, Melville, NY) mixed in saline. Cages of 3–5-day-old, starved, female Anopheles stephensi mosquitoes were then allowed to feed on individual mice for 30 minutes. Unfed mosquitoes were removed immediately after exposure, and remaining mosquitoes maintained at 24°C and a relative humidity of 80%. Eight to twelve days post-feeding, mosquito midguts were removed, stained with 0.1% mercurochrome (Sigma, St. Louis, MO), and malaria oocysts were enumerated. Sporozoite burden was determined on day 14 in 2 experiments by homogenizing pooled mosquito thoraxes, at least 50 per group, in approximately 1 mL of Hanks’ balanced salt solution supplemented with 1% normal mouse serum. The homogenate was then filtered through nylon mesh, re-suspended to a volume of 4 mL, and aliquots examined for sporozoites by hemocytometer. The resulting pooled sporozoite burden was expressed as the number of sporozoites per mosquito.

Statistical analysis.

Normally distributed variables were compared using Student’s unpaired t-test. Oocyst counts were compared using Student’s unpaired t-test after log10 transformation. Comparison of proportions was performed using the chi-square test. Bivariate analysis was performed by calculating Pearson’s correlation coefficient between the geometric mean number of oocysts per mosquito for each cage of mosquitoes fed on individual mice and the following terms: anemia, P. yoelii asexual parasitemia, and P. yoelii gametocytemia at the time of feeding. Analysis was performed using Microsoft (Redmond, WA) Excel® and STATA version 8.1 (STATA Corporation, College Station, TX).

RESULTS

Increased infectivity and parasite burden in mosquitoes fed on co-infected mice.

To determine whether chronic helminth co-infection modulated the intensity of P. yoelii transmission, An. stephensi mosquitoes were fed on E. caproniP. yoelii co-infected or P. yoelii only–infected mice at day five post-malaria infection. Over four replicate experiments, mosquitoes exposed to co-infected mice had a greater proportion of oocyst-positive mosquitoes (80.1%, n = 241) than those exposed to malaria only–infected mice (72.0%, n = 232, χ2 = 4.27, P = 0.039; Table 1).

We next compared midgut parasite burden of infected mosquitoes between groups. As shown in Figure 1 for pooled data over the four replicates, mosquitoes fed on co-infected mice had a significantly greater geometric mean number of oocysts (19.2 oocysts per mosquito, n = 167) than those exposed to P. yoelii only–infected mice (10.5 oocysts, n = 193, P = 0.004). Differences between groups were also significant for pooled data, as well as for three of the four individual replicates (P = 0.034, n ≥ 65 per group; P = 0.002, n ≥ 50 per group; P = 0.002, n ≥ 30 per group; P = 0.820, n ≥ 25 per group).

In two of the four experiments, we also compared the resulting sporozoite burden between groups. In one experiment, mosquitoes fed on co-infected mice had approximately 10-fold more sporozoites per mosquito than those fed on malaria only–infected controls (7,150 versus 750 sporozoites). In the other experiment, an approximately two-fold increase (2,120 versus 1,260 sporozoites; n ≥ 50 mosquitoes per group).

Gametocyte and total parasite levels in mice were not significantly different between co-infected and malaria only–infected mice on the day of mosquito feeding day five post-malaria infection (P = 0.60 and P = 0.08, respectively; Table 2). We also found no correlation between gametocytemia of individual mice and the numbers of oocysts detected in mosquitoes fed on either co-infected mice or malaria only–infected mice (r = −0.44 and −0.38, respectively). Likewise, there was no correlation between total parasitemia and oocyst burden for either group (r = −0.07 and = 0.57, respectively).

Additionally, packed cell volumes were similar between groups at day five of malaria infection (P = 0.11), and also did not correlate with oocyst burdens for either co-infected mice or malaria only–infected mice (r = 0.53 and = 0.75, respectively).

Temporal restriction of P. yoelii infectivity.

Successful transmission of P. yoelii rapidly decreases at day six after an initial peak of maximal infectivity between days two and five.28 To determine whether chronic helminth co-infection mitigated this temporal restriction of transmission, mosquitoes were exposed to E. caproni–malaria co-infected or malaria only–infected mice on days 8 and 13 of malaria infection in each of 2 separate experiments. Although gametocytes and asexual malaria parasites were present in both groups of mice at similar levels on days 8 or 13 of malaria infection (Table 2), no oocysts were detected in mosquitoes fed on either group of mice (n ≥ 100 mosquitoes per group, combined). Helminth infection therefore seemed to have no effect on abrogating the temporal restriction of P. yoelii transmission.

DISCUSSION

Co-infection with various helminth species occurs commonly in malaria-endemic areas, and the potential for helminths to influence malaria transmission has not been closely examined. In the present study, using a mouse model of E. caproni and P. yoelii co-infection, we observed that concurrent intestinal helminth infection in mice resulted in a significantly increased proportion of malaria-infected mosquitoes and a significantly increased parasite burden in infected mosquitoes at both oocyst and salivary gland sporozoite levels. We also found that the restriction of P. yoelii transmission to the first five days of infection was not extended by helminth co-infection.

Co-infecting parasites have previously been shown to influence transmission of mosquito-borne pathogens. Filarial larvae and malaria sporozoites were found to facilitate increased arbovirus dissemination in mosquitoes by disrupting the mosquito midgut and salivary glands, respectively.3036 In another study,37 filarial co-infection had no effect on malaria parasite transmission from humans to naturally fed mosquitoes, although low intensity of malaria infection may have precluded detection of an effect. Such mechanical disruptions are not possible in the present E. caproniP. yoelii system because echinostome parasites exclusively inhabit the gut and do not produce blood stage forms capable of being ingested with malaria parasites during mosquito feeding.

The central findings of these studies were the consistent increase in the proportion of Plasmodium-infected mosquitoes and increased intensity of parasite burden in mosquitoes fed on co-infected mice over those fed on malaria only–infected controls. This consistency is remarkable given the inherent biologic variability in these types of experiments. So that these trends may be fully appreciated, we have presented the primary data by experiment in addition to pooled results. Increased mosquito infectivity and parasite burden could not be attributed to differences in gametocyte or asexual malaria parasite density or anemia between groups of mice. The similarity of gametocytemia contrasts with a previous report of increased P. falciparum gametocyte prevalence in helminth co-infected humans, although this increase was nullified after adjustment for anemia.29 Furthermore, we also did not observe a correlation between gametocytemia and oocyst production in either co-infected mice or malaria-only infected controls. A lack of correlation between gametocytemia and oocyst burden is routinely noted,3841 and is likely due to the predilection for rodent malaria gametocytes to sequester in capillaries close to the skin.4244 Consequently, even if differences in gametocyte prevalence were detected between groups, it would be difficult to prove increased parasite transmission based solely on peripheral gametocyte density. Gametocyte density in the present study varied greatly between mice, and development of a P. yoelii gametocyte-specific quantitative detection assay, similar to the one recently designed for P. chabaudi,45 would prove useful in further investigating the relationship between gametocyte density and malaria transmission to mosquitoes.

Anemia is a prominent feature of many human helminth infections,46,47 and represents a significant risk factor for malaria gametocyte carriage.48 In the present study, which used E. caproni, a non-hematophagous intestinal parasite,49 no significant difference was detected in packed cell volume between co-infected mice and those infected only with malaria (Table 2). Although prediuresis, the process by which mosquitoes concentrate erythrocytes during blood feeding, may compensate for low levels of anemia,50 the non-significant difference in anemia detected in the present study is unlikely to account for the observed increase in oocyst burden. One would predict that mosquitoes fed on anemic co-infected mice would have ingested fewer gametocyte-infected erythrocytes than those fed on malaria only-infected controls. We did not compare packed cell volume between groups beyond day five because we had already determined that transmission did not occur at later time points.

Echinostoma caproni co-infection increases the intensity and duration of late stage P. yoelii infection,22 yet in the present study, we found that it did not alter the temporal restriction of P. yoelii transmission. On the basis of studies in P. berghei, decreases in host pH and bicarbonate levels induced by increasing parasitemia seem to prevent exflagellation of microgametes independently of other serum-derived transmission modulating factors.7 Because P. berghei and P. yoelii share similar patterns of transmission restriction, it is likely that host physiologic changes also influence infectivity of P. yoelii parasites. We conclude therefore that although E. caproni infection alters the magnitude of transmission during early infection, it does not abrogate the ensuing physiologic alterations that limit P. yoelii transmission temporally. This restriction, however, is unique to rodent malaria parasites, and the possibility remains for helminths to alter transmission throughout the course of patent, and even sub-patent, human malaria infections.

In summary, we have shown that infection in the vertebrate host with an intestinal helminth can enhance transmission of malaria parasites to the mosquito vector. We hypothesize that such interactions are likely due to helminth-induced changes to the host immune system. Because similar enhancement of malaria transmission may also occur during widespread helminth co-infection in humans, current global deworming strategies may realize benefits beyond the alleviation of helminth infections. However, in light of studies that found an increase in malaria infections after anthelmintic treatment,19,51 further studies are required to properly evaluate such predicted outcomes and the mechanisms mediating enhanced transmission.

Table 1

Mosquito infectivity in four experiments in Anopheles stephensi mosquitoes fed on either Echinostoma caproni–Plasmodium yoelii co-infected (E + M) or P. yoelii only–infected (M) mice five days post-malaria infection

Oocyst-positive mosquitoes/Total no (%)
ExperimentE + MM
* P = 0.039, by χ2 test.
171/78 (91.0)62/73 (84.9)
256/69 (81.2)49/71 (69.0)
334/43 (79.1)28/42 (66.7)
432/51 (62.8)28/46 (60.9)
Total193/241 (80.1)167/232 (72.0)*
Table 2

Plasmodium yoelii gametocytemia, parasitemia, and packed cell volume (PCV) in Echinostoma caproni–P. yoelii co-infected (E + M), or P. yoelii-only–infected (M) BALB/c mice at indicated days post-malaria infection*

E + MM
Mean ± SDMean ± SDNP
* ND = not done.
† By Student’s t-test.
Gemotocytemia (%)
    Day 50.032 ± 0.0320.026 ± 0.018≥ 130.60
    Day 80.015 ± 0.0140.020 ± 0.013≥ 100.46
    Day 130.128 ± 0.1430.098 ± 0.050≥ 40.71
Total parasitemia (%)
    Day 54.0 ± 1.82.9 ± 1.4≥ 130.08
    Day 85.8 ± 2.94.8 ± 3.3≥ 100.74
    Day 1337.5 ± 12.522.2 ± 13.1≥ 40.12
PCV (%)
    Day 552.2 ± 5.056.8 ± 4.160.11
    Day 8NDND0
    Day 13NDND0
Figure 1.
Figure 1.

Box plots of number of Plasmodium yoelii oocysts per mosquito in oocyst-positive Anopheles stephensi fed on Echinostoma caproniP. yoelii co-infected mice (E+M) (n = 193 mosquitoes) or P. yoelii only–infected mice (M) (n = 167 mosquitoes) five days post-malaria infection (aggregate of four experiments). Boxes, with medians indicated, contain the middle 50% of observations (the interquartile range [IQR]), whiskers indicate the smallest and largest observations within the lower and upper fence (± 1.5 × IQR), and outliers are observations outside the upper fence. Geometric mean number of oocysts were 19.2 (E+M) and 10.5 (M) (P = 0.004, by Student’s t-test of log-transformed values.)

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 76, 6; 10.4269/ajtmh.2007.76.1052

*

Address correspondence to Nirbhay Kumar, Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, 615 North Wolfe Street, Baltimore, MD 21205, Telephone: 410-955-7177, Fax: 410-955-0105. E-mail: nkumar@jhsph.edu

Authors’ addresses: Gregory S. Noland and Nirbhay Kumar, Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, 615 North Wolfe Street, Baltimore, MD 21205, Telephone: 410-955-7177, Fax: 410-955-0105, E-mails: gnoland@jhsph.edu and nkumar@jhsph.edu. Thaddeus K. Graczyk, Departments of Environmental Health Sciences and Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, 615 North Wolfe Street, Baltimore, MD 21205 Telephone: 410-614-4984, Fax: 410-955-0105, E-mail: tgraczyk@jhsph.edu. Bernard Fried, Department of Biology, Lafayette College, Kunkel Hall 204, Easton, PA 18042, Telephone: 610-330-5463, Fax: 610-330-5705, E-mail: friedb@lafayette.edu.

Acknowledgments: We thank Erik J. Fitzgerald for statistical advice.

Financial support: Gregory S. Noland is supported by a predoctoral fellowship from the Johns Hopkins Malaria Research Institute. Research in the laboratory of Nirbhay Kumar is supported by grants from the National Institutes of Health.

REFERENCES

  • 1

    Karunaweera ND, Carter R, Grau GE, Kwiatkowski D, del Giudice G, Mendis KN, 1992. Tumour necrosis factor-dependent parasite-killing effects during paroxysms in non-immune Plasmodium vivax malaria patients. Clin Exp Immunol 88 :499–505.

    • Search Google Scholar
    • Export Citation
  • 2

    Naotunne TS, Karunaweera ND, Del Giudice G, Kularatne MU, Grau GE, Carter R, Mendis KN, 1991. Cytokines kill malaria parasites during infection crisis: extracellular complementary factors are essential. J Exp Med 173 :523–529.

    • Search Google Scholar
    • Export Citation
  • 3

    Kaushal DC, Carter R, Rener J, Grotendorst CA, Miller LH, Howard RJ, 1983. Monoclonal antibodies against surface determinants on gametes of Plasmodium gallinaceum block transmission of malaria parasites to mosquitoes. J Immunol 131 :2557–2562.

    • Search Google Scholar
    • Export Citation
  • 4

    Rener J, Graves PM, Carter R, Williams JL, Burkot TR, 1983. Target antigens of transmission-blocking immunity on gametes of Plasmodium falciparum.J Exp Med 158 :976–981.

    • Search Google Scholar
    • Export Citation
  • 5

    Naotunne TD, Rathnayake KD, Jayasinghe A, Carter R, Mendis KN, 1990. Plasmodium cynomolgi: serum-mediated blocking and enhancement of infectivity to mosquitoes during infections in the natural host, Macaca sinica.Exp Parasitol 71 :305–313.

    • Search Google Scholar
    • Export Citation
  • 6

    Peiris JS, Premawansa S, Ranawaka MB, Udagama PV, Munasinghe YD, Nanayakkara MV, Gamage CP, Carter R, David PH, Mendis KN, 1988. Monoclonal and polyclonal antibodies both block and enhance transmission of human Plasmodium vivax malaria. Am J Trop Med Hyg 39 :26–32.

    • Search Google Scholar
    • Export Citation
  • 7

    Butcher GA, Sinden RE, Billker O, 1996. Plasmodium berghei: infectivity of mice to Anopheles stephensi mosquitoes. Exp Parasitol 84 :371–379.

    • Search Google Scholar
    • Export Citation
  • 8

    de Silva NR, Brooker S, Hotez PJ, Montresor A, Engels D, Savioli L, 2003. Soil-transmitted helminth infections: updating the global picture. Trends Parasitol 19 :547–551.

    • Search Google Scholar
    • Export Citation
  • 9

    Mwangi TW, Bethony JM, Brooker S, 2006. Malaria and helminth interactions in humans: an epidemiological viewpoint. Ann Trop Med Parasitol 100 :551–570.

    • Search Google Scholar
    • Export Citation
  • 10

    Buck AA, Anderson RI, MacRae AA, 1978. Epidemiology of poly-parasitism. IV. combined effects on the state of health. Trop Med Parasitol 29 :253–268.

    • Search Google Scholar
    • Export Citation
  • 11

    Nacher M, Singhasivanon P, Yimsamran S, Manibunyong W, Thanyavanich N, Wuthisen R, Looareesuwan S, 2002. Intestinal helminth infections are associated with increased incidence of Plasmodium falciparum malaria in Thailand. J Parasitol 88 :55–58.

    • Search Google Scholar
    • Export Citation
  • 12

    Spiegel A, Tall A, Raphenon G, Trape JF, Druilhe P, 2003. Increased frequency of malaria attacks in subjects co-infected by intestinal worms and Plasmodium falciparum malaria. Trans R Soc Trop Med Hyg 97 :198–199.

    • Search Google Scholar
    • Export Citation
  • 13

    Le Hesran JY, Akiana J, Ndiaye EH, Dia M, Senghor P, Konate L, 2004. Severe malaria attack is associated with high prevalence of Ascaris lumbricoides infection among children in rural Senegal. Trans R Soc Trop Med Hyg 98 :397–399.

    • Search Google Scholar
    • Export Citation
  • 14

    Sokhna C, Le Hesran JY, Mbaye PA, Akiana J, Camara P, Diop M, Ly A, Druilhe P, 2004. Increase of malaria attacks among children presenting concomitant infection by Schistosoma mansoni in Senegal. Malar J 3 :43.

    • Search Google Scholar
    • Export Citation
  • 15

    Nacher M, Gay F, Singhasivanon P, Krudsood S, Treeprasertsuk S, Mazier D, Vouldoukis I, Looareesuwan S, 2000. Ascaris lumbricoides infection is associated with protection from cerebral malaria. Parasite Immunol 22 :107–113.

    • Search Google Scholar
    • Export Citation
  • 16

    Nacher M, Singhasivanon P, Silachamroon U, Treeprasertsuk S, Vannaphan S, Traore B, Gay F, Looareesuwan S, 2001. Helminth infections are associated with protection from malaria-related acute renal failure and jaundice in Thailand. Am J Trop Med Hyg 65 :834–836.

    • Search Google Scholar
    • Export Citation
  • 17

    Lyke KE, Dicko A, Dabo A, Sangare L, Kone A, Coulibaly D, Guindo A, Traore K, Daou M, Diarra I, Sztein MB, Plowe CV, Doumbo OK, 2005. Association of Schistosoma haematobium infection with protection against acute Plasmodium falciparum malaria in Malian children. Am J Trop Med Hyg 73 :1124–1130.

    • Search Google Scholar
    • Export Citation
  • 18

    Briand V, Watier L, Hesran JY, Garcia A, Cot M, 2005. Coin-fection with Plasmodium falciparum and Schistosoma haematobium: protective effect of schistosomiasis on malaria in Senegalese children? Am J Trop Med Hyg 72 :702–707.

    • Search Google Scholar
    • Export Citation
  • 19

    Brutus L, Watier L, Briand V, Hanitrasoamampionona V, Razanatsoarilala H, Cot M, 2006. Parasitic co-infections: Does As-caris lumbricoides protect against Plasmodium falciparum infection? Am J Trop Med Hyg 75 :194–198.

    • Search Google Scholar
    • Export Citation
  • 20

    Lwin M, Last C, Targett GA, Doenhoff MJ, 1982. Infection of mice concurrently with Schistosoma mansoni and rodent malarias: contrasting effects of patent S. mansoni infections on Plasmodium chabaudi, P. yoelii and P. berghei.Ann Trop Med Parasitol 76 :265–273.

    • Search Google Scholar
    • Export Citation
  • 21

    Christensen NØ, Furu P, Kurtzhals J, Odaibo A, 1988. Heterologous synergistic interactions in concurrent experimental infection in the mouse with Schistosoma mansoni, Echinostoma revolutum, Plasmodium yoelii, Babesia microti, and Trypanosoma brucei.Parasitol Res 74 :544–551.

    • Search Google Scholar
    • Export Citation
  • 22

    Noland GS, Graczyk TK, Fried B, Fitzgerald EJ, Kumar N, 2005. Exacerbation of Plasmodium yoelii malaria in Echinostoma caproni infected mice and abatement through anthelmintic treatment. J Parasitol 91 :944–948.

    • Search Google Scholar
    • Export Citation
  • 23

    Helmby H, Kullberg M, Troye-Blomberg M, 1998. Altered immune responses in mice with concomitant Schistosoma mansoni and Plasmodium chabaudi infections. Infect Immun 66 :5167–5174.

    • Search Google Scholar
    • Export Citation
  • 24

    Yoshida A, Maruyama H, Kumagai T, Amano T, Kobayashi F, Zhang M, Himeno K, Ohta N, 2000. Schistosoma mansoni infection cancels the susceptibility to Plasmodium chabaudi through induction of type 1 immune responses in A/J mice. Int Immunol 12 :1117–1125.

    • Search Google Scholar
    • Export Citation
  • 25

    Su Z, Segura M, Morgan K, Loredo-Osti JC, Stevenson MM, 2005. Impairment of protective immunity to blood-stage malaria by concurrent nematode infection. Infect Immun 73 :3531–3539.

    • Search Google Scholar
    • Export Citation
  • 26

    Yan Y, Inuo G, Akao N, Tsukidate S, Fujita K, 1997. Down-regulation of murine susceptibility to cerebral malaria by inoculation with third-stage larvae of the filarial nematode Brugia pahangi.Parasitology 114 :333–338.

    • Search Google Scholar
    • Export Citation
  • 27

    Graham AL, Lamb TJ, Read AF, Allen JE, 2005. Malaria-filaria coinfection in mice makes malarial disease more severe unless filarial infection achieves patency. J Infect Dis 191 :410–421.

    • Search Google Scholar
    • Export Citation
  • 28

    Bastien P, Landau I, Baccam D, 1987. Inhibition of the infectivity of Plasmodium gametocytes by the serum of the parasite host. Perfecting an experimental model. Ann Parasitol Hum Comp 62 :195–208.

    • Search Google Scholar
    • Export Citation
  • 29

    Nacher M, Singhasivanon P, Silachamroon U, Treeprasertsu S, Krudsood S, Gay F, Mazier D, Looareesuwan S, 2001. Association of helminth infections with increased gametocyte carriage during mild falciparum malaria in Thailand. Am J Trop Med Hyg 65 :644–647.

    • Search Google Scholar
    • Export Citation
  • 30

    Mellor PS, Boorman J, 1980. Multiplication of bluetongue virus in Culicoides nubeculosus (Meigen) simultaneously infected with the virus and the microfilariae of Onchocerca cervicalis (Railliet & Henry). Ann Trop Med Parasitol 74 :463–469.

    • Search Google Scholar
    • Export Citation
  • 31

    Paulson SL, Poirier SJ, Grimstad PR, Craig GB Jr, 1992. Vector competence of Aedes hendersoni (Diptera: Culicidae) for la crosse virus: Lack of impaired function in virus-infected salivary glands and enhanced virus transmission by sporozoite-infected mosquitoes. J Med Entomol 29 :483–488.

    • Search Google Scholar
    • Export Citation
  • 32

    Turell MJ, Mather TN, Spielman A, Bailey CL, 1987. Increased dissemination of dengue 2 virus in Aedes aegypti associated with concurrent ingestion of microfilariae of Brugia malayi.Am J Trop Med Hyg 37 :197–201.

    • Search Google Scholar
    • Export Citation
  • 33

    Vaughan JA, Turell MJ, 1996. Dual host infections: enhanced infectivity of eastern equine encephalitis virus to Aedes mosquitoes mediated by Brugia microfilariae. Am J Trop Med Hyg 54 :105–109.

    • Search Google Scholar
    • Export Citation
  • 34

    Vaughan JA, Turell MJ, 1996. Facilitation of rift valley fever virus transmission by Plasmodium berghei sporozoites in Anopheles stephensi mosquitoes. Am J Trop Med Hyg 55 :407–409.

    • Search Google Scholar
    • Export Citation
  • 35

    Vaughan JA, Trpis M, Turell MJ, 1999. Brugia malayi microfilariae (Nematoda: Filaridae) enhance the infectivity of Venezuelan equine encephalitis virus to Aedes mosquitoes (Diptera: Culicidae). J Med Entomol 36 :758–763.

    • Search Google Scholar
    • Export Citation
  • 36

    Zytoon EM, el Belbasi HI, Matsumura T, 1993. Mechanism of increased dissemination of Chikungunya virus in Aedes albopictus mosquitoes concurrently ingesting microfilariae of Dirofilaria immitis.Am J Trop Med Hyg 49 :201–207.

    • Search Google Scholar
    • Export Citation
  • 37

    Burkot TR, Molineaux L, Graves PM, Paru R, Battistutta D, Dagoro H, Barnes A, Wirtz RA, Garner P, 1990. The prevalence of naturally acquired multiple infections of Wuchereria bancrofti and human malarias in anophelines. Parasitology 100 :369–375.

    • Search Google Scholar
    • Export Citation
  • 38

    Munderloh UG, Kurtti TJ, 1987. The infectivity and purification of cultured Plasmodium berghei ookinetes. J Parasitol 73 :919–923.

  • 39

    Ponnudurai T, Lensen AH, Van Gemert GJ, Bensink MP, Bolmer M, Meuwissen JH, 1989. Infectivity of cultured Plasmodium falciparum gametocytes to mosquitoes. Parasitology 98 :165–173.

    • Search Google Scholar
    • Export Citation
  • 40

    Haji H, Smith T, Charlwood JD, Meuwissen JH, 1996. Absence of relationships between selected human factors and natural infectivity of Plasmodium falciparum to mosquitoes in an area of high transmission. Parasitology 113 :425–431.

    • Search Google Scholar
    • Export Citation
  • 41

    Sinden RE, Butcher GA, Billker O, Fleck SL, 1996. Regulation of infectivity of Plasmodium to the mosquito vector. Adv Parasitol 38 :53–117.

    • Search Google Scholar
    • Export Citation
  • 42

    Gautret P, Gantier JC, Baccam D, Miltgen F, Saulai M, Chabaud AG, Landau I, 1996. The gametocytes of Plasmodium vinckei petteri, their morphological stages, periodicity and infectivity. Int J Parasitol 26 :1095–1101.

    • Search Google Scholar
    • Export Citation
  • 43

    Gautret P, Miltgen F, Chabaud AG, Landau I, 1996. Synchronized Plasmodium yoelii yoelii: pattern of gametocyte production, sequestration and infectivity. Parassitologia 38 :575–577.

    • Search Google Scholar
    • Export Citation
  • 44

    Gautret P, Miltgen F, Gantier JC, Chabaud AG, Landau I, 1996. Enhanced gametocyte formation by Plasmodium chabaudi in immature erythrocytes: pattern of production, sequestration, and infectivity to mosquitoes. J Parasitol 82 :900–906.

    • Search Google Scholar
    • Export Citation
  • 45

    Wargo AR, Randle N, Chan BH, Thompson J, Read AF, Babiker HA, 2006. Plasmodium chabaudi: reverse transcription PCR for the detection and quantification of transmission stage malaria parasites. Exp Parasitol 112 :13–20.

    • Search Google Scholar
    • Export Citation
  • 46

    Hotez PJ, Brooker S, Bethony JM, Bottazzi ME, Loukas A, Xiao S, 2004. Hookworm infection. N Engl J Med 351 :799–807.

  • 47

    Friedman JF, Kanzaria HK, McGarvey ST, 2005. Human schistosomiasis and anemia: The relationship and potential mechanisms. Trends Parasitol 21 :386–392.

    • Search Google Scholar
    • Export Citation
  • 48

    Price R, Nosten F, Simpson JA, Luxemburger C, Phaipun L, ter Kuile F, van Vugt M, Chongsuphajaisiddhi T, White NJ, 1999. Risk factors for gametocyte carriage in uncomplicated falciparum malaria. Am J Trop Med Hyg 60 :1019–1023.

    • Search Google Scholar
    • Export Citation
  • 49

    Wisnewski N, Fried B, Halton DW, 1986. Growth and feeding of Echinostoma revolutum on the chick chorioallantois and in the domestic chick. J Parasitol 72 :684–689.

    • Search Google Scholar
    • Export Citation
  • 50

    Taylor PJ, Hurd H, 2001. The influence of host haematocrit on the blood feeding success of Anopheles stephensi: implications for enhanced malaria transmission. Parasitology 122 :491–496.

    • Search Google Scholar
    • Export Citation
  • 51

    Murray J, Murray A, Murray M, Murray C, 1978. The biological suppression of malaria: an ecological and nutritional interrelationship of a host and two parasites. Am J Clin Nutr 31 :1363–1366.

    • Search Google Scholar
    • Export Citation
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