Molecular Characterization of Leishmania Parasites in Giemsa-Stained Slides from Cases of Human Cutaneous and Visceral Leishmaniasis,Eastern Algeria

Abstract

Background:

In Algeria, visceral leishmaniasis (VL) is due to Leishmania (L.) infantum, while three cutaneous forms (CL) are caused by Leishmania major, Leishmania tropica and Leishmania infantum. In this study, the use of Giemsa-stained slides was evaluated with two PCR techniques, in Eastern Algeria.

Materials and Methods:

A total of 136 samples corresponding to 100 CL smears (skin scrapings) and 36 VL slides (bone marrow aspirates) collected from 2008 to 2014 were tested. Upon DNA extraction, two PCRs were used to amplify the ribosomal Internal Transcribed Spacer 1 (ITS1) and mini-exon genes. Amplified products were digested (PCR-RFLP) and profiles analyzed for Leishmania species identification. A statistical analysis was also performed.

Results:

ITS1-PCR was found significantly more sensitive than mini-exon-PCR (77.95% positives vs. 67.65%; p = 0.001). Comparison of PCR positivity showed statistically significant differences between old and recently prepared slides suggesting a better use of recent slides in PCR analyses. For species identification, PCR-restriction fragment length polymorphism (RFLP) results of ITS1 and mini-exon were concordant. L. infantum was identified from VL cases and L. infantum, L. major, and L. tropica from CL ones. According to geographical origin, L. infantum was found in North-Eastern provinces, while L. major was distributed from the North to the Center-East of Algeria. Interestingly, two L. tropica samples were identified in Annaba, located far North-East Algeria.

Conclusion:

Distribution of leishmaniasis in Eastern parts of Algeria, besides finding of L. tropica in the far North, is in this study described for the first time using molecular tools, thus confirming the usefulness of slides for PCR identification of Leishmania parasites in retrospective epidemiological investigations.

Introduction

Leishmaniasis is a group of zoonotic diseases that infect the viscera (visceral leishmaniasis: VL), the skin (cutaneous leishmaniasis: CL), and mucous membranes (mucocutaneous leishmaniasis: MCL). They are caused by intracellular parasites of the genus Leishmania (L.) and are transmitted by bites of phlebotomine sand flies. VL and CL are endemic in Algeria. Indeed, around 44,000 cases of CL and 111 cases of VL are reported annually which makes this country 1 of the 10 countries with the highest estimated cases of CL (Alvar et al. 2012). VL in Algeria is due to Leishmania infantum in the northern parts of the country (Belazzoug et al. 1985). However, during the last few years, it has extended from the North to the South with several sporadic cases (Benallal et al. 2013, Adel et al. 2014).

CL is caused by three species: L. infantum in the North, Leishmania major in central and southern parts of the country, and Leishmania killicki, which is a variant of Leishmania tropica, with sporadic cases in Ghardaïa province (Harrat et al. 2009, Boubidi et al. 2011).

The difficulties on the diagnosis of leishmaniasis are due to the low number of parasites found in clinical samples, the similar morphologies of Leishmania species, and the ambiguity of symptoms. Thus, direct detection of parasites embedded within slide smears is always performed by clinicians to establish disease diagnosis. Other techniques are also used to detect the parasites, like in vitro culture and serology. However, they have several disadvantages (Srivastava et al. 2011). Identification of the parasite species can only be done using molecular techniques. Indeed, the identification of Leishmania species based on their geographical origin or clinical signs and symptoms can be problematic because multiple species of Leishmania coexist in endemic regions, as well as several species can cause both visceral and cutaneous disease (Adel et al. 2014). Molecular tools like PCR become especially useful in such situations.

PCR and PCR-restriction fragment length polymorphism (RFLP) techniques targeting different genes and sequences from the kinetoplast and nuclear Leishmania genomes have been extensively used for their detection and identification (Guerbouj et al. 2014). Using PCR, biological samples have been used for diagnosis of leishmaniasis, such as bone marrow, blood, and lymph nodes (Piarroux et al. 1994, Lachaud et al. 2001, Cortes et al. 2004). Skin and mucosal biopsies and buffy coat were also used (Pirmez et al. 1999, Lachaud et al. 2001). Moreover, PCR has made possible the use of dried or old material. Indeed, it was possible to extract and amplify Leishmania DNA from Giemsa-stained slides (Motazedian et al. 2002, Al-Jawabreh et al. 2006, Volpini et al. 2006, Brustoloni et al. 2007), paraffin-embedded tissues (Lanús et al. 2005, de Lima et al. 2011), and blood or bone marrow aspirates spotted on filter paper (El Tai et al. 2000, Cortes et al. 2004, Da Silva et al. 2004).

This study aimed to evaluate the use of PCR in leishmaniasis diagnosis using Giemsa-stained slides from bone marrow aspirates or skin scrapings prepared during routine diagnosis of leishmaniasis by microscopy, in Eastern Algeria. Performance of two PCR techniques as retrospective diagnostic tools was assessed and causal parasite species were identified by PCR-RFLP analyses.

Materials and Methods

Clinical samples

The study included 136 slides corresponding to 100 CL smears from skin scrapings and 36 from bone marrow aspirate of leishmaniasis patients that were referred to the Parasitology/Mycology laboratory of the University hospital of Annaba, North East Algeria, during the 2008–2014 periods to establish diagnosis of leishmaniasis. For each sample, information concerning the patient's clinical manifestations, geographical origin of infection (Fig. 1), age, and sex was recorded (Supplementary Table S1; Supplementary Data are available online at www.liebertpub.com/vbz). It is worth noting that for all patients, travel history to endemic leishmaniasis regions was taken into account for determining the geographical origin of infection. For diagnosis purposes, slides were MGG (May Grünwald Giemsa) stained for microscopic testing (observation of Leishmania amastigotes) and stored at room temperature for further molecular testing.

FIG. 1.

Geographical distribution of leishmaniasis patients collected in Eastern provinces of Algeria and the identified causal Leishmania species. The study area is shown within a square in the map of Algeria to the upper left corner of each panel. (A, B) Represent the geographical origin of VL and CL cases, respectively. Gray coding shows the number of cases collected from each province. Species identification made by ITS1 and mini-exon PCR-restriction fragment length polymorphism (RFLP) is shown by pie charts, where size is proportional to the number of identified cases. CL, cutaneous leishmaniasis; ITS1, ribosomal Internal Transcribed Spacer 1; VL, visceral leishmaniasis.

Microscopy observation of slides

Slides were scanned for amastigotes under 100 × oil-immersion light microscopy. Size of the scanned area corresponded to an area of ∼1000 oil-immersion fields (OIF), as 1000 OIF per slide must be screened before declaring negative (Al-Jawabreh et al. 2006, World Health Organization 2010). The average amastigote density is graded following the number of parasites observed per field (viewed with a 10× eyepiece and 100× oil-immersion lens).

Control parasites

Tunisian strains from the L. infantum (MHOM/TN/94/LV49 and MHOM/TN/94/LV50) and L. major (MPSM/TN/87/RON114) species and a strain from Iraq representing L. tropica (MHOM/IQ/76/Bag9) were used in all molecular analyses as positive controls. These strains were obtained after in vitro culture and were characterized using isoenzyme typing and ribosomal Internal Transcribed Spacer 1 (ITS1) PCR-RFLP (Bensoussan et al. 2006).

DNA extraction

To perform DNA extraction, each smear was scraped off slides using a sterile scalpel. A lysis buffer (50 mM Tris-HCl pH7.4, 10 mM EDTA, 50 mM NaCl) was added to the scrapped material and incubated overnight at 55°C with 100 μg/mL Proteinase K and 0.05% SDS. Total DNA was phenol/chloroform purified and ethanol precipitated (Guizani et al. 1994). Also using phenol/chloroform method, DNAs were extracted from in vitro cultured parasites representing L. major, L. infantum, and L. tropica species that were used as positive controls.

PCR and PCR-RFLP analysis

Two PCR protocols were applied to the DNAs extracted from the slides. The first PCR targeted a Leishmania-specific ITS1 gene that was amplified using the primers LITS and L5.8S to generate PCR products in the range of 300 to 350 bp, as previously described (Bensoussan et al. 2006). The second PCR amplified mini-exon genes using primers Fme and Rme and generated 400 to 450 bp fragments, as shown previously (Marfurt et al. 2003b). Primers sequences used in the different PCRs, reaction conditions, and cycling are detailed in Supplementary Table S2.

To perform PCR-RFLP analysis, ITS1 and mini-exon PCR products were further digested using HaeIII (Vivantis, Malaysia) and EaeI (Biolabs, France) restriction enzymes, respectively, as previously described (Marfurt et al. 2003a, Bensoussan et al. 2006, Serin et al. 2007). All reactions were incubated overnight at 37°C. Resulting restriction fragments were separated on 3% agarose gels and fragment sizes were estimated by comparison with bands of 100 bp (Vivantis) and 50 bp (Promega, France) DNA size ladders. Control Leishmania strains (L. major, L. infantum, and L. tropica) were also PCR-RFLP analyzed and run in parallel, thus constituting the necessary controls of the study (Fig. 2).

FIG. 2.

Identification of Leishmania species by ITS1 PCR-HaeIII RFLP and Mini-exon PCR-EaeI RFLP. (A, B) Illustrate examples of PCR-RFLP results. (A) Shows HaeIII restriction analysis of ITS1-PCR products and (B) shows EaeI restriction analysis of mini-exon PCR products. Lanes where sample codes are shown corresponding to CL or VL cases that were identified as Leishmania infantum species. Lanes 1 to 4 correspond to CL cases that were identified as Leishmania major (1, 3, and 4) and Leishmania tropica (2). Codes of the latter cases are 1: 215/13; 2: 39/12; 3: 80/12, and 4: 136/12. Leishmania species representative strains are L. infantum (MHOM/TN/94/LV49), L. major (MPSM/TN/87/RON114), and L. tropica (MHOM/IQ/76/Bag9). Sizes presented on the left were extracted from Schönian et al. (2003), Marfurt et al. (2003b), Bensoussan et al. (2006), and Serin et al. (2007). All sizes are indicated in bp. M is 50 bp molecular weight marker.

Statistical analysis

Collected data are summarized using frequencies and relative frequencies (percentages). Between-group comparisons were carried out using chi-square test or Fisher exact test for variables expressed in frequencies. Slides used in this study were collected according to the cases available over 7 years, from 2008 to 2014. However, because the number of analyzed slides drastically differed between corresponding years of collection, they were classified into two periods, those collected recently (years 2012 to 2014) and the older ones collected from 2008 to 2011. PCR positive results obtained in the two periods were cumulated and compared using the nonparametric Wilcoxon signed-rank test for paired data. For all analyses, statistically significant differences were those with a p value <0.05. Statistical analysis was performed using SPSS (version 17.0 for Windows).

Results

PCR amplification of Leishmania DNA from slides

Comparing sensitivity of both PCR targets showed ITS1 PCR to be significantly more sensitive than mini-exon PCR (p < 0.05). Indeed, over a total of 136 DNAs extracted from slides made from CL (skin scrapings) and VL (bone marrow aspirates) patients, ITS1, and mini-exon amplified fragments at the expected size were obtained in the case of 106 (77.95%) and 92 (67.65%) samples, respectively (Table 1). Moreover, 90 DNAs showed positive signals with both ITS1 and mini-exon PCRs. However, 16 more slides were amplified only with ITS1 PCR, and two cases were amplified with mini-exon PCR only (Table 1).

Table 1.

Comparison of Internal Transcribed Spacer 1 and Mini-Exon PCR assay performance

	CL and VL samples			CL samples			VL samples
	Mini-exon PCR			Mini-exon PCR			Mini-exon PCR
	Positive results	Negative results	Total (%)	Positive results	Negative results	Total (%)	Positive results	Negative results	Total (%)
ITS1 PCR
Positive results	90	16	106 (77.95)	70	13	83 (83)	20	3	23 (63.9)
Negative results	2	28	30 (22.05)	2	15	17 (17)	0	13	13 (36.1)
Total (%)	92 (67.65)	44 (32.3)	136 (100)	72 (72)	28 (28)	100 (100)	20 (55.56)	16 (44.44)	36 (100)
p ^a	0.001			0.007			NS

p Values were calculated with chi-squared tests following ITS1 and mini-exon PCR results' comparisons using cross-tabulation between positive and negative results of both PCRs.

CL, cutaneous leishmaniasis; ITS1, ribosomal Internal Transcribed Spacer 1; NS, nonsignificant; VL, visceral leishmaniasis.

As regards the CL group, ITS1 PCR was also found to be significantly more sensitive than mini-exon PCR (Table 1), while this difference was not found significant in the VL group, which is constituted with a fewer number of samples than the CL group (100 vs. 36). This could explain the nonsignificance of the statistical analysis (Table 1).

The positive slides for Leishmania amastigotes were classified as “very abundant” (VA or +++; one or more parasites are found in 1 to 10 fields), “abundant” (A or ++; one or more parasites are observed in 10 to 100 fields), and “poor” (P or +; one or more parasites are found in 100 to 1000 fields). Richness of slides with amastigote forms was compared to results of the two PCRs used and was shown to be in agreement with them. Accordingly, slides that were classified as amastigote A and VA showed very few negative ITS1 or mini-exon PCR results (no amplification). Indeed, ITS1 PCR showed no amplification in only 2 out of the 22 (9.1%) A slides and in 2 out of the 42 (4.8%) VA slides, while 26 negative results were found over 72 (36.1%) amastigote P slides (Table 2). The mini-exon PCR showed negative results in 3 (13.64%) and 5 (11.9%) cases of A and VA slides, respectively, while in P slides 36 cases (50%) could not be amplified. These differences were shown to be statistically significant (p < 0.005, Table 2).

Table 2.

PCR Results According to Richness of Slides

	ITS1 PCR			Mini-exon PCR
	(P) ^a	(A) ^a	(VA) ^a	(P) ^a	(A) ^a	(VA) ^a
Positive results (%)	46 (63.89)	20 (90.9)	40 (95.2)	36 (50)	19 (86.36)	37 (88.1)
Negative results (%)	26 (36.11)	2 (9.1)	2 (4.8)	36 (50)	3 (13.64)	5 (11.9)
Total	72	22	42	72	22	42
p ^b	<0.0001			<0.0001

Richness of slides with amastigote Leishmania forms: (P) (poor or +), one or more parasites are found in 100 to 1000 fields; (A) (abundant or ++), one or more parasites are found in 10 to 100 fields; and (VA) (very abundant or +++), one or more parasites are found in 1 to 10 fields.

p Values were calculated with chi-squared tests following between-group comparisons according to richness of slides (P, A, and VA), using cross-tabulation between positive and negative results of both PCRs.

Correlation between PCR positivity and oldness of slides showed amplification results of ITS1 and mini-exon PCRs reaching 84.28% and 75.72%, respectively, in recent slides collected in 2012–2014 period (Table 3). However, for the older slides (2008–2011 period), percentage of positive ITS1 and mini-exon PCR results decreased to 71.21% and 59.1%, respectively (Table 3). These differences were found to be statistically significant when using mini-exon PCR (p < 0.05) and at a border of significance when using ITS1 PCR (p = 0.066, Table 3), which overall showed that more positive PCR results would be found when using recently prepared slides than older ones.

Table 3.

Correlation Between PCR Positivity and Oldness of Slides

	Years
	2008	2009	2010	2011	Total: 2008–2011 ^a	2012	2013	2014	Total: 2012–2014 ^a	p ^b
ITS1 PCR^c (%)	5/9 (55.56)	8/13 (61.54)	13/17 (76.47)	21/27 (77.78)	47/66 (71.21)	15/24 (62.5)	34/35 (97.15)	10/11 (90.91)	59/70 (84.28)	NS
Mini-exon PCR^c (%)	5/9 (55.56)	4/13 (30.77)	12/17 (70.59)	18/27 (66.67)	39/66 (59.1)	11/24 (45.83)	33/35 (94.29)	9/11 (81.82)	53/70 (75.72)	0.038

Cumulative positive results obtained in the period of years indicated.

p Values were calculated following between-group of years' period (indicated in b) comparisons.

Number of PCR positive slides over the total number of analyzed slides.

NS, nonsignificant.

Leishmania species identification by PCR-RFLP analysis

PCR-RFLP analysis provided discriminating species-specific RFLP profiles that allowed to precisely identify Leishmania species within slides. However, it is worth noting that some of the PCR-RFLP profiles generated could not be interpreted (Supplementary Fig. S1). This was specifically the case of 9 and 10 CL slides, using ITS1 and mini-exon PCR-RFLPs, respectively, which were eliminated from the RFLP analysis.

Restriction profiles of ITS1 and mini-exon amplified fragments from VL slide DNAs identified L. infantum as the infecting species (Fig. 2). In contrast, the three species L. major, L. infantum, and L. tropica were identified from slides corresponding to CL samples (Fig. 2). Indeed, using either ITS1 or mini-exon PCR-RFLPs, L. infantum species was the predominant species (48/74 [64.9%] and 45/62 [72.6%], respectively) followed by L. major (24/74 [32.4%] and 15/62 [24.2%], respectively) and L. tropica (2/74 [2.7%] and 2/62 [3.2%], respectively) (Table 4). In addition, in all positive slides for both PCR targets, results of ITS1 PCR-HaeIII and mini-exon PCR-EaeI were concordant for the identification of Leishmania parasites. Thus, L. infantum was found using both tools in 20 VL cases, and 3 DNAs were identified by only ITS1 PCR-HaeIII RFLP (Table 4). For CL slides, results of both tools were concordant in case of 44 DNAs corresponding to L. infantum and 15 DNAs identified as L. major. The L. tropica species was found using both tools within two slides (Table 4).

Table 4.

Leishmania Species Identification by PCR-RFLP Analyses

		ITS1 and Mini-exon RFLPs ^a	ITS1-HaeIII ^b	Mini-exon-EaeI ^c	Total ^d
VL samples	Number of successfully PCR-RFLP analyzed slides^e/Total number of analyzed slides^f		23/23	20/20
	Leishmania Infantum	20	3	0	23
CL samples	Number of successfully PCR-RFLP analyzed slides^e/Total number of analyzed slides^f		74/83	62/72
	L. infantum	44	4	1	49
	Leishmania major	15	9	0	24
	Leishmania tropica	2	0	0	2

Number of cases where the corresponding species was identified by both tools.

Number of cases where the corresponding species was identified by ITS1-HaeIII but not with mini-exon-EaeI.

Number of cases where the corresponding species was identified by mini-exon-EaeI but not with ITS1-HaeIII.

The total number of the corresponding species identified by ITS1-HaeIII and/or mini-exon-EaeI.

The number of successfully PCR-RFLP analyzed slides is the number of cases where PCR-RFLP profiles were interpretable without ambiguity.

Total number of analyzed slides is the number of PCRs positive cases that were further digested with the corresponding restriction enzymes.

PCR-RFLP, polymerase chain reaction-restriction fragment length polymorphism.

Geographical distribution of identified species

Giemsa-stained slides that were used in this study were in most cases from leishmaniasis patients that originated from three provinces situated in Northeastern Algeria (Annaba, Souk Ahras, and Guelma). Distribution of identified species according to likely geographical origin of infection showed presence of L. infantum VL in Northern provinces of Algeria especially in Souk Ahras where 12 cases were recorded. VL cases were also sporadically distributed in the Center, with one case in Biskra province. Cutaneous cases due to L. infantum were found located in the North (region of Guelma, Annaba, and Souk Ahras) while those due to L. major were distributed from the north to the Center-Eastern provinces. Interestingly, L. tropica was identified in two CL patients and was mapped in Annaba, located far Northeast Algeria (Fig. 1 and Supplementary Table S3).

Discussion

Identification of Leishmania species by microscopic examination of Giemsa-stained slides is not possible since Leishmania species are morphologically similar (Schönian et al. 2003). In the past decade, molecular techniques using mainly PCR have been used for the diagnosis of leishmaniasis (Tavares et al. 2011). In endemic regions where more than one Leishmania species is present, diagnostic tools are required for detection and identification of parasites directly within samples (Schönian et al. 2003).

Giemsa-stained bone marrow aspirates or skin scrapings on glass slides, prepared during routine microscopy diagnosis of leishmaniasis, are valuable sources of DNA for PCR diagnosis (Schönian et al. 2003, Al-Jawabreh et al. 2006, Brustoloni et al. 2007). Indeed, slides can be easily stored at room temperature and used to perform DNA extraction and PCR amplification. Specifically, in the case of VL patients, PCR diagnosis using DNA extracted from Giemsa-stained bone marrow slides is a suitable method to confirm diagnosis (Brustoloni et al. 2007, Pandey et al. 2013). This can be helpful when reevaluating the diagnosis of controversial cases or in retrospective epidemiological studies. It is also a very useful tool for the diagnosis of difficult cases, having a low number of parasite load and negative microscopy results (Brustoloni et al. 2007, Pandey et al. 2013).

Our study demonstrates value of Giemsa-stained slides not only for leishmaniasis diagnosis but also for retrospective epidemiological studies, in Algeria. In total 136 slides corresponding to CL smears and bone marrow aspirates of leishmaniasis patients were studied using ITS1 and mini-exon PCR targets. ITS1 amplified fragments at the expected sizes were obtained in 77.95% of the cases, distributed as 83% CL and 63.9% VL samples. Previous studies using Giemsa-stained slides for amplification with ITS1 PCR showed that Leishmania DNA was detected in 96.2% of slides representing suspected CL cases in Morocco (Arroub et al. 2013). In Libya, 195 out of 450 CL samples (43.33%) were successfully amplified with ITS1 PCR (Amro et al. 2012).

PCR targeting mini-exon genes was found statistically less sensitive than ITS1 PCR. Gene copy numbers could probably explain these differences since this criterion has been previously shown to be crucial for better amplification results, especially when using poor quality DNA (Schönian et al. 2003). To our knowledge, this is the first work reporting statistical comparison of ITS1 and mini-exon PCRs for diagnosis of leishmaniasis from stored slides. Sensitivity differences could also be explained by variations at the level of amplification efficacy in relation with parasite burden, presence of inhibitors from host material, or reaction conditions. Indeed, in this work, using both PCR targets we found positivity of PCRs to be significantly related to the richness with amastigote forms which can partly explain that more positive results were found in CL slides than in VL ones as it is known that parasite number is more elevated in CL smears than in bone marrow aspirates (de Lima et al. 2011, Pandey et al. 2013). Previous studies showed sensitivity of PCR protocols to vary outstandingly when applied either on cultured parasites or on clinical samples (Schönian et al. 2003). In addition, different PCR sensitivity levels were found according to sources of clinical samples used (spleen, skin, liver, and bone marrow) (Schönian et al. 2003).

Investigation using statistical analysis of the oldness of slides in relation to the amplification results of ITS1 and mini-exon PCRs showed that more positive results would be found when using recently prepared slides than older ones. However, Leishmania DNA was successfully amplified from slides stored for up to 36 years in Brazil (Volpini et al. 2006). More recent studies showed amplification of Leishmania DNA from slides that were stored between 2 and 10 years (Pandey et al. 2013).

Upon amplification of Leishmania DNA from Giemsa-stained slides, it is important to precisely identify the corresponding species. Thus, PCR-RFLP analysis which generates discriminating species-specific profiles allowed identifying L. infantum as the infecting species from slides of VL cases, and the three species L. major, L. infantum, and L. tropica as causing CL cases. These findings confirm the occurrence of the three species in Algeria and give new insights into their distribution in eastern parts of the country. Indeed, VL cases are mainly distributed in Northern provinces; however, scarce VL cases were also found in the Center, as previously described, denoting a spread of VL to the central and southern parts of Algeria (Benallal et al. 2013, Adel et al. 2014). Cutaneous samples from which L. infantum was identified were mainly from provinces in the far North like previously known (Belazzoug et al. 1985), while two cases that originated from the East-Center (Tebessa), one case from the Center (M'sila), and one case from the South (El Oued) are in this study originally described. Besides, finding L. infantum species in the majority of CL cases of East Algeria is atypical for other regions of Northern Africa, specifically in Tunisia where L. major has been described as the most frequent species in CL (Aoun and Bouratbine 2014). This is mainly related to eco-epidemiological changes probably affecting the vector and reservoir ecology and behavior, thus influencing the geographical distribution of the disease and its causing Leishmania species (Toumi et al. 2012).

In this study, cases from which L. major species were identified originated from North and Center-Eastern provinces, thus confirming the spread of CL form into Northern parts of Algeria, as previously described using the MLEE technique (Boudrissa et al. 2012). Interestingly, L. tropica was identified in Annaba, far North-East Algeria. This is the first report describing the occurrence of this species in this part of the country looking at Giemsa-stained slides. Indeed, this species also known as L. killicki has been first described in Ghardaïa, South of Algeria (Harrat et al. 2009, Boubidi et al. 2011), then a few cases were described in Constantine region in the North (Boudrissa et al. 2012). Recently, two L. killicki cases identified by amplification and sequencing of the topoisomerase II gene from Giemsa-stained slides were found in Tipaza region located North-Center Algeria (Izri et al. 2014). In Tunisia, sporadic CL cases due to L. tropica have been first described in a microfocus in Tataouine, South-Eastern Tunisia and then various studies confirmed its presence and showed its spread to Central and South-Western Tunisia (Ben Abda et al. 2009, Bousslimi et al. 2010). However, this species has never been described in the far North of the country, as it is the case in Algeria. Overall, our results show the need to reevaluate the distribution of L. tropica infection in Algeria and further study its transmission cycle.

Conclusions

As described in this study, usefulness of PCR techniques on Giemsa-stained slides for retrospective epidemiological analysis of leishmaniasis in Eastern Algeria is shown. Parasite species identification assessed by ITS1 or mini-exon PCR-RFLP analyses confirmed L. infantum as responsible of VL and also CL cases. L. major and L. tropica were also found in CL samples, which give new insights into the actual distribution of ethiological agents of leishmaniases in Eastern parts of the country. The occurrence of L. tropica in Annaba, far Northeast Algeria, is in this study described for the first time within patients' samples using molecular tools. ITS1 PCR was found more sensitive than mini-exon PCR, amplifying DNA from slides up to 7 years old. Giemsa-stained slides constitute a valuable source of DNA that could be used to compare Leishmania transmitted years ago with recent ones using appropriate DNA assays.

Footnotes

Acknowledgments

This work was supported by the Ministry of Higher Education and Scientific Research in Tunisia (LR11IPT04) and the Ministry of Public Health and the Ministry of Higher Education and Scientific Research in Algeria FNR (DGRSDT) & ANDRS (ATRSS, Ex: ANDRS). S.G. and R.M. received a bilateral Tunisian-Algerian cooperation grant [EEP47_2013-15].

Author Disclosure Statement

No competing financial interests exist. The study was approved by the Ethics Committee of the ATRSS agency (Agence Thématique de Recherche en Sciences de la Santé), Ex-ANDRS agency (Agence Nationale pour le Développement de la Recherche en Santé) of the Ministry of Higher Education and Scientific Research in Algeria (reference number: 01/02/05/03/04/020), and the Ethics Committee of the “Badji Mokhtar, Annaba's University” in Algeria (reference number: 352/26/04/2015).

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