Leishmania – Microscopy and PCR

Consistent with O. Reg. 671/92 of the French Language Services Act, laboratory testing information on this page is only available in English because it is scientific or technical in nature and is for use only by qualified health care providers and not by members of the public.

This page provides routine microscopy and polymerase chain reaction (PCR) testing information for leishmaniasis at Public Health Ontario (PHO). The causative agent of leishmaniasis is the genus Leishmania. Manifestations include cutaneous leishmaniasis (CL), mucosal leishmaniasis (ML), and visceral leishmaniasis (VL, also known as kala azar).

This page is for information limited to microscopy and PCR. For serology, please refer to the following PHO webpage:

Testing Indications

CL and ML: PCR is the main diagnostic test method and should be requested whenever testing is indicated. Microscopic examination is helpful when positive but has poor sensitivity in CL and ML. Serology has limited value in CL and ML.

VL: Microscopy and PCR are the main diagnostic test methods and should be requested whenever testing is indicated. Serology is an adjunct but not a replacement for microscopy and PCR in VL.

Specimen Collection and Handling

Specimen Requirements

Test Requested Required Requisition(s) Specimen Type Minimum Volume Collection Kit

Leishmania, cutaneous leishmaniasis, or mucosal leishmaniasis

Lesion brushing/ swabbing


Lesion scraping


Lesion aspirate


Filter paper lesion impression


Lesion biopsy





0.1 ml sterile saline


Filter paper


~2-4 mm punch biopsy

Sterile container

Leishmania, visceral leishmaniasis, or kala azar

Whole blood in EDTA

Splenic, bone marrow, or lymph node aspirate

Bone marrow, lymph node, or liver biopsy

4.0 ml


1.0 ml



EDTA tube


EDTA tube or sterile container


Sterile container

(any type)

Formalin-fixed paraffin-embedded (FFPE) tissue blocks with accompanying histology stained slides


FFPE block and slides(s)

Submission and Collection Notes


See Special Instructions below for suggestions on specimen collection for CL and ML.

Complete all fields of the requisition form, including:
  • Test(s) requests and indications for testing
  • Patient setting/population
  • Clinical information including symptom onset date or if asymptomatic
  • Travel history – the country/region of exposure MUST be stated otherwise testing may be delayed or cancelled.

To assist in testing prioritization, state in the requisition if the patient is pregnant, has HIV, has impaired immunity, or is in the intensive care unit.


Label the specimen container(s) with the patient’s first and last name, date of collection, and one other unique identifier such as the patient’s date of birth or Health Card Number. Failure to provide this information may result in rejection or testing delay.

Storage and Transport

Place specimen in biohazard bag and seal. Specimens should be stored at room temperature following collection and shipped to PHO’s laboratory as soon as possible. All clinical specimens must be shipped in accordance to the Transportation of Dangerous Good Act.

Special Instructions

CL or ML: To increase the likelihood of detection, collect multiple specimens from the most active areas (e.g. oozing ulcer base and advancing inflammatory margins) of each lesion using different techniques as below. Avoid healed areas or areas with secondary bacterial/fungal infection. Clean areas with 70% alcohol (not iodine) and remove overlying necrotic debris/scabs. If needed, a low concentration of local anesthetics (e.g. 1% lidocaine) can be used to reduce discomfort.

    1. If using lesion scraping technique: scrape most active areas (without making incision; bleeding should be minimal but oozing may occur) and transfer scraped material into a sterile container with minimal amount of sterile saline to keep material moistened during transit.
    2. If using lesion brushing/swabbing technique: brush most active areas multiple times until pink-tinged tissue fluids are wicked onto brush/swab and transfer brush/swab into a sterile container with minimal amount of sterile saline keep material moistened during transit.
    3. If using lesion aspirate technique: using 0.1-0.2 ml of sterile saline and a 23- to 27-gauge needle, gently insert and rotate the needle back and forth tangentially in the most active area while aspirating pink-tinged tissue fluids up to the needle hub (note: if insufficient fluid is obtained, inject 0.05-0.1 ml of sterile saline in the area and resume suction) and transfer aspirated fluid into a sterile container.
    4. If using filter paper lesion impression technique: using sterile coarse-porosity 7-cm filter paper, gently press paper onto ulcer base multiple times to cover ulcer surface until pink-tinged tissue fluids are wicked onto the paper and transfer paper into a sterile container.
    5. If using lesion biopsy technique: obtain a 2-4 mm punch biopsy of the most active area (down to the level of the dermis) and transfer biopsied material into a sterile container with minimal amount of sterile saline to keep material moistened during transit.

Note that PCR sensitivity varies by cutaneous or mucosal lesion type, lesion severity, collection technique, lesion area sampled, and Leishmania species. Boggild et al.1,2 reported comparable sensitivity between filter paper impressions (86.6-92.3%) and aspirates or scrapings (92.5-94.2%). Valencia et al.3 similarly reported comparable sensitivity between filter paper impressions (99.1%), foam swabs (99.1%), and scrapings (96.6%). Boggild et al.4 reported comparable sensitivity between foam swabs (95.7%) and biopsy specimens (95.7%). Adams et al.5 reported superior sensitivity of foam swabs (97.5%) versus aspirates (80%). Blaizot et al.6 reported superior sensitivity of post-biopsy cotton swabs (98%) versus biopsy specimens (89%). Suarez et al.7 reported superior parasite loads at the ulcer base compared to the raised border, as well as superior parasite loads using foam swabs or scrapings versus biopsies.

VL: PCR sensitivity also varies by specimen type and Leishmania species. Although splenic aspirates are usually considered of highest yield, their collection is a high-risk procedure, and blood and bone marrow specimens alternatively also have high accuracy (93.1-95.3% sensitivity and 92.6-95.6% specificity per de Ruiter et al.8).

Requisitions and Kit Ordering

Test Frequency and Turnaround Time (TAT)

Leishmania PCR and microscopy are performed Monday to Friday at Public Health Ontario’s Toronto laboratory site.

Turnaround time is up to 2 days for PCR and 3 days for microscopy from receipt at PHO’s laboratory.

If positive, samples are forwarded to the US CDC for species identification. Turnaround time for species identification is up to 3 weeks.

Test Methods

Leishmania PCR detection is conducted at PHO using three different laboratory-developed assays performed in parallel: Wortmann et al.9 Leishmania spp. 18S real-time PCR, Mary et al.10 Leishmania (Leishmania)spp. kDNA real-time PCR, and Lopez et al.11 Leishmania (Viannia)spp.end-point PCR.

Leishmania microscopy is conducted at PHO using Giemsa staining. Whole blood specimens undergo buffy coat concentration.

Leishmania species identification is conducted at the US CDC using ITS1 SYBR Green PCR melt curve analysis (de Almeida et al.)12 and ITS2 Sanger sequencing (de Almeida et al.)13.


PCR is performed on all specimens received. If PCR is positive, the specimen will be forwarded to the US CDC for Leishmania species identification.

Microscopy is only performed on specimens if requested on the requisition.


The following table provides possible PCR test results with associated interpretations:

Leishmania spp. 18S target (Wortmann)

Leishmania (Leishmania) kDNA target (Mary)

Leishmania (Viannia) kDNA target (Lopez)


Positive or Negative



Leishmania spp. detected. Species identification to follow.




Leishmania spp. detected. Probable Leishmania subgenus. Species identification to follow.


Negative or Positive (low level)


Leishmania spp. detected. Probable Viannia subgenus. Species identification to follow.










Leishmania spp. indeterminate. Testing of additional specimens recommended if clinically indicated.





Leishmania spp. not detected. Multiple specimens are required to rule out leishmaniasis.




Invalid due to failed detection of the assay internal controls. May be due to inhibitors in the specimen. Testing of additional specimens recommended if clinically indicated.


The following table provides microscopy test results with associated interpretation:

Parasite microscopy


Amastigotes detected.

Unable to distinguish Leishmania from Trypanosoma cruzi amastigotes by microscopy.

No amastigotes detected.

Negative microscopy results do not rule out leishmaniasis. PCR has superior sensitivity over microscopy.


Test Performance:
At PHO, microscopy has an overall sensitivity of 27% and specificity of 100% (Lau et al.14). The Leishmania (Leishmania) kDNA real-time PCR has a very high sensitivity for the Leishmania subgenus with about 40% cross-reactivity with the Viannia subgenus. The Leishmania (Viannia) kDNA end-point PCR has very high sensitivity for the Viannia subgenus with minimal cross-reactivity with the Leishmania subgenus (Cruz et al.15). The species identification assays may be unable to provide identification at low parasite loads. Refer to Special Instructions above for further details on specimen type performance and limitations.


Results are reported to the physician, authorized health care provider (General O. Reg 45/22, s.18) or submitter as indicated on the requisition.


  1. Boggild AK, Valencia BM, Espinosa D, Veland N, Ramos AP, Arevalo J, et al. Detection and species identification of Leishmania DNA from filter paper lesion impressions for patients with American cutaneous leishmaniasis. Clin Infect Dis. 2010;50(1):e1-6. Available from: https://doi.org/10.1086/648730
  2. Boggild AK, Ramos AP, Valencia BM, Veland N, Calderon F, Arevalo J, et al. Diagnostic performance of filter paper lesion impression PCR for secondarily infected ulcers and nonulcerative lesions caused by cutaneous leishmaniasis. J Clin Microbiol. 2011;49(3):1097-100. Available from: https://doi.org/10.1128/JCM.02457-10
  3. Valencia BM, Veland N, Alba M, Adaui V, Arevalo J, Low DE, et al. Non-invasive cytology brush PCR for the diagnosis and causative species identification of American cutaneous leishmaniasis in Peru. PLoS One. 2012;7(11):e49738. Available from: https://doi.org/10.1371/journal.pone.0049738
  4. Boggild AK, Valencia BM, Veland N, Pilar Ramos A, Calderon F, Arevalo J, et al. Non-invasive cytology brush PCR diagnostic testing in mucosal leishmaniasis: superior performance to conventional biopsy with histopathology. PLoS One. 2011;6(10):e26395. Available from: https://doi.org/10.1371/journal.pone.0026395
  5. Adams ER, Gomez MA, Scheske L, Rios R, Marquez R, Cossio A, et al. Sensitive diagnosis of cutaneous leishmaniasis by lesion swab sampling coupled to qPCR. Parasitology. 2014;141(14):1891-7. Available from: https://doi.org/10.1017/S0031182014001280
  6. Blaizot R, Simon S, Ginouves M, Prévot G, Blanchet D, Ravel C, et al. Validation of swab sampling and SYBR green-based real-time PCR for the diagnosis of cutaneous leishmaniasis in French Guiana. J Clin Microbiol. 2021;59(2):e02218-20. Available from: https://doi.org/10.1128/JCM.02218-20
  7. Suárez M, Valencia BM, Jara M, Alba M, Boggild AK, Dujardin JC, et al. Quantification of Leishmania (Viannia) kinetoplast DNA in ulcers of cutaneous leishmaniasis reveals inter-site and inter-sampling variability in parasite load. PLoS Negl Trop Dis. 2015;9(7):e0003936. Available from: https://doi.org/10.1371/journal.pntd.0003936
  8. de Ruiter CM, van der Veer C, Leeflang MM, Deborggraeve S, Lucas C, Adams ER. Molecular tools for diagnosis of visceral leishmaniasis: systematic review and meta-analysis of diagnostic test accuracy. J Clin Microbiol. 2014;52(9):3147-55. Available from: https://doi.org/10.1128/JCM.00372-14
  9. Wortmann G, Sweeney C, Houng HS, Aronson N, Stiteler J, Jackson J, et al. Rapid diagnosis of leishmaniasis by fluorogenic polymerase chain reaction. Am J Trop Med Hyg. 2001;65(5):583-7. Available from: https://doi.org/10.4269/ajtmh.2001.65.583
  10. Mary C, Faraut F, Lascombe L, Dumon H. Quantification of Leishmania infantum DNA by a real-time PCR assay with high sensitivity. J Clin Microbiol. 2004;42(11):5249-55. Available from: https://doi.org/10.1128/JCM.42.11.5249-5255.2004
  11. Lopez M, Inga R, Cangalaya M, Echevarria J, Llanos-Cuentas A, Orrego C, et al. Diagnosis of Leishmania using the polymerase chain reaction: a simplified procedure for field work. Am J Trop Med Hyg. 1993;49(3):348-56. Available from: https://doi.org/10.4269/ajtmh.1993.49.348
  12. de Almeida ME, Koru O, Steurer F, Herwaldt BL, da Silva AJ. Detection and differentiation of Leishmania spp. in clinical specimens by use of a SYBR green-based real-time PCR assay. J Clin Microbiol. 2016;55(1):281-90. Available from: https://doi.org/10.1128/JCM.01764-16
  13. de Almeida ME, Steurer FJ, Koru O, Herwaldt BL, Pieniazek NJ, da Silva AJ. Identification of Leishmania spp. by molecular amplification and DNA sequencing analysis of a fragment of rRNA internal transcribed spacer 2. J Clin Microbiol. 2011;49(9):3143-9. Available from: https://doi.org/10.1128/JCM.01177-11   
  14. Lau R, Valencia B, Ralevski F, Llanos-Cuentas A, Boggild AK. Validation of a molecular diagnostic algorithm for leishmania detection in a non-endemic setting. Int J Infect Dis. 2014;21(1):162-3. Available from: https://doi.org/10.1016/j.ijid.2014.03.761
  15. Cruz I, Millet A, Carrillo E, Chenik M, Salotra P, Verma S, et al. An approach for interlaboratory comparison of conventional and real-time PCR assays for diagnosis of human leishmaniasis. Exp Parasitol. 2013;134(3):281-9. Available from: https://doi.org/10.1016/j.exppara.2013.03.026   
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Updated 16 Aug 2023