Diagnostic Accuracy of Polymerase Chain Reaction Against Giemsa Staining on Tissue Biopsy for Cutaneous Leishmaniasis

By Aleena Khalid, Maria Mushtaq Gill, Irfan Ali Mirza, Sakeenah Hussain Naqvi, Syed Adeel Hussain Gardezi, Rabia Sajjad

Affiliations

1. Department of Microbiology, Armed Forces Institute of Pathology / National University of Medical Sciences, Rawalpindi, Pakistan

doi: 10.29271/jcpsp.2024.09.1024

ABSTRACT
Objective: To evaluate the diagnostic accuracy of a commercial real-time polymerase chain reaction (PCR) kit targeting 18S rRNA against Giemsa-stained tissue slides in patients clinically suspected of cutaneous leishmaniasis (CL).
Study Design: Cross-sectional analytical study.
Place and Duration of the Study: Department of Microbiology, Armed Forces Institute of Pathology / National University of Medical Sciences, Rawalpindi, Pakistan, from July to December 2022.
Methodology: Samples of skin tissue in 98 patients suspected of CL were evaluated. These samples were subjected to Giemsa-staining for microscopy and real-time PCR. Sensitivity, specificity, and accuracy of the PCR were calculated keeping Giemsa-stained tissue slide microscopy as gold standard.
Results: Out of the 98 tissue samples, 37 were found positive for leishmaniasis on PCR while 13 were found Leishmania positive on microscopy of Giemsa-stained slides. The sensitivity, specificity, and accuracy of the PCR for the detection of Leishmania species were 100%, 71.8%, and 91.8%, respectively with 100% negative predictive value.
Conclusion: This study demonstrates that the commercial PCR is a reliable diagnostic test for the diagnosis of CL. The ease, rapidity, and reliability of the PCR make it a dependable tool in diagnostic repertoire of CL.

Key Words: Giemsa stain, Leishmania spp., Polymerase chain reaction, Viasure.

INTRODUCTION

Leishmaniasis is the third most significant global vector-borne disease which spreads via infected sandfly bites.^1,2 There are over 20 Leishmania species causing human infections.¹ This infection, found in approximately 88 countries, predominantly thrives in regions characterised by tropical and subtropical climates. Among these regions, Afghanistan, Brazil, Iran, Saudi Arabia, Peru, and Syria stand out with over 90% of reported infections originating from these nations.³ It manifests in various forms including visceral (VL), cutaneous (CL) and mucosal (ML). Cutaneous leishmaniasis (CL) is the most prevalent among them, accounting for approximately 0.6 to 1 million new cases reported annually.⁴
 

CL manifests as a skin disorder characterised by the appearance of one or more lesions, typically presenting as ulcerated or verrucous plaques, on parts of the body within the reach of the vector sandflies. In some cases, the infection may extend to involve the lymph nodes and satellite lesions may also develop, albeit infrequently.⁵

Cutaneous form of leishmaniasis is more widespread in Pakistan.⁶ In the 2013-2015 epidemic, war-affected Waziristan saw a 3.61% prevalence of CL.⁷ A cross-sectional study conducted across four Khyber Pakhtunkhwa districts including Karak, Lakki Marwat, Tank, and Dera Ismail Khan revealed a 50.4% prevalence of CL among participants with skin lesions.⁸ Over the last decade, the country has experienced an anticipated annual incidence of 4.6 cases per 1,000 individuals.⁹

CL is also known as the great imitator, as it can mimic various dermatoses. Such similarity occasionally leads to misdiagnosis, resulting in inappropriate treatment and health issues.¹⁰ This demands early and accurate diagnosis to replace costly and complex treatment regimens with substantial adverse effects and increasing resistance concerns.¹¹ The gold standard for the diagnosis of CL is the parasite-based tests including microscopy, parasite isolation by in vitro culture or inoculation of mice and hamsters.¹² The immunological tests including skin-prick tests and ELISA for antibody detection are also available with limited reliability.

Currently, molecular techniques for diagnosis of leishmaniasis are being increasingly adopted because of their rapidity.^11,13 Among these methods, the real-time PCR gives ease of usage, rapidity of result, real-time result analysis, and low-risk of contamination. A commercial real-time PCR kit targeting 18S rRNA claims to be highly sensitive and specific for the diagnosis of leishmaniasis. However, it has never been evaluated in Pakistan for its diagnostic sensitivity claims.⁴ Therefore, this study was planned to evaluate the diagnostic accuracy of this commercial real-time PCR Giemsa-stained tissue slide microscopy in patients clinically suspected of CL.

METHODOLOGY

This was a cross-sectional analytical study, conducted at the Department of Microbiology, Armed Forces Institute of Pathology / National University of Medical Sciences, Rawalpindi, Pakistan. The sample size was calculated by using Buderer’s formula.¹⁴ Considering sensitivity of real-time PCR to be 93.9%, specificity as 100%, prevalence of Leishmania spp. as 50.4%, confidence level 95%, and margin of error 5%. The minimum sample size for this study was calculated to be 98.⁸

The sampling method employed in this study was non-probability convenience sampling, wherein individuals were conveniently selected based on their availability and suitability for the research. Patients of all ages and both genders, presenting with documented non-healing ulcers persisting for one month or more, and seeking medical attention at a tertiary care hospital in Rawalpindi, were eligible for inclusion. However, patients who had received partial or complete treatment for CL were excluded from the study population. Prior to participation, individuals provided informed consent and completed a detailed data collection form. Dermatologist then meticulously collected skin tissue biopsy specimens from the margins of the ulcers in a sterile manner which were subsequently dispatched to the study place for further analysis and examination.

These tissue fragments were finely ground using a scalpel and petri dish. Each sample was then divided into two parts. The initial step involved subjecting the first tissue fragment to Giemsa staining for microscopy. Films were made from tissue fragments, dried in the air then fixed by immersing in methanol for 10 minutes. Then the fixed films were transferred to a staining jar containing May-Grunwald stain freshly diluted with an equal volume of buffered water. After the films were allowed to stain for about 15 minutes, they were transferred without washing to a jar containing Giemsa stain freshly diluted with 9 volumes of buffered water. After staining for fifteen minutes, they were transferred to a jar containing buffered water and rapidly washed in four changes of water and left to stand for five minutes.¹⁵ After air drying, the stained slides were examined using a light microscope, employing oil immersion, with the aim of identifying any presence of Leishmania amastigotes within the tissue samples. The presence of Leishmania amastigotes can be seen in positive samples as shown in Figure 1.

Figure 1: Giemsa-stained slide demonstrating amastigotes taken under X1000 oil immersion lens.

The subsequent segment of the ground tissue specimen underwent real-time PCR analysis. To initiate DNA extraction, the Merck Millipore's Bacterial Xpress Nucleic Acid Extraction Kit was employed.¹⁶ To extract DNA, approximately 200 μl of extraction reagent, along with 50 μl of the sample, and an additional 10 μl of internal standard were combined within a microcentrifuge tube and thoroughly mixed via vortexing. Following a five-minute incubation period at room temperature, 250 μl of isopropyl alcohol was introduced into each tube, vigorously vortexed, and subsequently subjected to centrifugation at 16,000g for 10 minutes. Upon completion of centrifugation, the supernatant was carefully discarded and the resulting pellet was subjected to a washing step with 400 μl of 70% ethanol, followed by another round of vortexing and centrifugation at 6,000g for 10 minutes. After the removal of the supernatant, the pellet was air-dried and subsequently re-suspended in 50 μl of water. For the subsequent PCR analysis, a Leishmania real-time PCR Detection kit was employed.¹⁷ In each Smart Cycler tube, 10 μl of the DNA extract was combined with 20 μl of the master mix. These prepared tubes were then positioned within the Bio-Rad CFX96™ System RT PCR machine and the amplification process was initiated following the protocol outlined in the kit instructions. It is crucial to note that any sample exhibiting a cycle threshold (Ct) value of less than 40 was deemed positive for the presence of Leishmania spp.⁴ The positive and negative results were recorded for each sample. Figure 2 shows a positive test.

The data analysis was conducted utilising the Statistical Package for the Social Sciences (SPSS) version 23. For qualitative variables, frequencies and percentages were computed, while for quantitative variables, the mean and standard deviation were calculated. Inferential statistics were employed to assess the diagnostic accuracy of PCR, utilising Giemsa-stained tissue slide microscopy as the gold standard. Sensitivity, specificity, positive predictive value (PPV), negative predictive value (NPV), and accuracy measures were determined to evaluate the performance of the PCR assay.¹² Cohen Kappa was calculated to estimate the level of agreement between the two tests.

Figure 2: The curves of PCR as interpreted by the system made in a positive sample. Green curve indicates the sample that came out to be positive. Blue curve indicates positive internal control.

RESULTS

Out of the total 98 subjects, the majority were males, constituting 95.9% (n = 94), while female patients accounted for 4.1% (n = 4), with an overall average age of 37.51 ± 12 years. Majority of patients belonged to Rawalpindi (66.30%) followed by Waziristan (23.50%).

A comparison between PCR results and Giemsa-stained slides revealed that out of the ninety-eight samples assessed, thirty-seven samples tested positive via PCR, whereas only thirteen samples were positive through Giemsa staining. The sensitivity of the commercial PCR kit for Leishmania spp. detection was calculated as 100%, with a specificity of 71.8%. The PPV was determined to be 54.7%, while the NPV was 100%. The overall diagnostic accuracy of the PCR was found to be 91.8%.

Considering that the positivity of CL via Giemsa-stained slide microscopy was lower compared to the index test (PCR), Staquet’s correction was applied to ascertain the sensitivity and specificity of the PCR under conditions where the reference standard was imperfect.¹⁸

The sensitivity of the Giemsa-stained specimen was taken to be 37% and specificity to be 100%.¹⁹ Therefore, post-application of Staquet’s correction, the ultimate sensitivity of the PCR kit in this study was determined to be 100%, while its specificity was calculated at 97%. Additionally, the Cohen Kappa value, assessing the agreement between the PCR and Giemsa-stained slides, was found to be 0.403, with a significant p-value of <0.001.

DISCUSSION

CL stands as the most overlooked tropical ailment worldwide as it is undiagnosed or is misdiagnosed.¹⁰ The present approaches to treating leishmaniasis lack considerable efficacy and frequently lead to notable adverse effects.¹¹ Leishmaniasis causes significant morbidity and mortality in low and middle-income countries (LIMCs).⁵ Swift and accurate diagnosis, along with the identification of specific species, plays a vital role in halting the advancement of leishmaniasis.¹³ The literature indicates that Giemsa-stained specimen microscopy demonstrates high specificity (100%) but its sensitivity is considerably low (37%).¹⁹ This study also indicated the same through the Cohen’s Kappa which suggests that both PCR and Giemsa-stained tissue specimens detect CL in a fairly consistent way but there is still room for improvement in the gold standard.

The tested PCR is designed for diagnosis of the genus Leishmania targeting 18s rRNA.⁴ It identifies the presence of Leishmania species but does not differentiate between the specific types. The findings indicated that the tested PCR had a sensitivity of 100%, specificity of 97%, PPV of 54.5%, NPV of 100%, and diagnostic accuracy of 91.8%. In literature, a study was conducted by Arnau et al., in which they evaluated the diagnostic accuracy of the presently studied PCR against Leishmania culture and another commercial PCR kit.⁴ The specificity in this study is 100%, which is comparable to this study (97%) but sensitivity was found to be much lower (81.8%) as compared to 100%. Likewise, the PPV and NPV were 100% and 72.7% which are quite different from this study.⁴ The reason for this difference in results is most likely because of the different reference methods in the two studies.

The gender distribution in this study was approximately 96% males and 4% females which is in contrast to another study conducted in the North Waziristan Agency.²⁰ In this study, they elaborated the pattern of CL in the North Waziristan agency and found that among microscopically confirmed CL patients, 57% were males while 43% were females.²⁰ The reason for the difference in gender distribution in both studies is likely because of the difference of patient catchment area.

Comparing the Giemsa-staining results of this study with those of a study by Nateghi et al., where the authors assessed two types of PCR alongside Giemsa-stained specimens, it becomes apparent that their Giemsa positivity rate was 77.27% (17/22), while this study indicates it to be 35% (13/37).²¹ This shows a significant disparity in the positivity rates between the two studies. The observed difference in positivity rates could be attributed to several factors, including unequal distributions of parasites within the lesion. It is plausible that the portion of the lesion collected for smear preparation may not have contained enough parasites for detection by microscopy, contributing to the lower positivity rate. Additionally, variation in the expertise of the microscopists involved in the examination of the stained slides could have influenced the detection and interpretation of parasites. Overall, these findings underscore the importance of considering various factors, such as lesion characteristics and expertise levels, when interpreting Giemsa staining results for the diagnosis of CL.

Among the patients included in the study, 80.6% (79/98) presented with solitary lesions, with or without crusting or purulent discharge, while 19.4% (19/98) had multiple lesions. Interestingly, this distribution contrasts with findings from a study conducted in Iraq by Ali et al., where multiple ulcers were more prevalent on the body (60.44%) compared to solitary ulcers (39.66%).²² One potential explanation for this disparity could be related to the demographic characteristics of the study’s patient population. Military recruits were primarily selected, and it is plausible that their health is monitored regularly, allowing for prompt observation and treatment of initial lesions before the development of additional ones. This proactive approach to healthcare may have led to a higher proportion of solitary lesions observed as compared to the Iraqi study.

Despite the accuracy of the tested PCR for diagnosis of CL, the high cost is still a challenge in resource-limited settings, however, the ease of use, rapid turnaround time, and consistent reliability make it a valuable addition to the diagnostic arsenal for this condition, facilitating timely intervention and treatment strategies. For implementation in such resource-limited settings, Giemsa-staining should be the first-line diagnostic tool, however, for patients with chronic skin lesions in areas of high endemicity, PCR can be used as a second-line confirmatory test for definitive diagnosis.

This study highlights that molecular detection using the tested PCR can be used for accurate and timely diagnosis of CL thus preventing the spread of the disease within the country. Based upon the increasing incidence of the disease and the associated morbidity, the association of the clinical picture with the species can be the future avenues of research. Further studies involving species identification and their association with disease activity and treatment response are recommended.

CONCLUSION

This study provides compelling evidence that PCR targeting 18S rRNA is a dependable diagnostic tool for the accurate diagnosis of CL.

ETHICAL  APPROVAL:
Ethical approval was taken by Institutional Review Board and Ethical Committee (Letter Number: FC-MIC21-2/READ-IRB/22/ 1914, Dated: 27 Jun 2022).

PATIENTS’  CONSENT:
Consent was obtained from all patients prior to their participation in the study.

COMPETING  INTEREST:
The authors declared no conflict of interest.

AUTHORS’  CONTRIBUTION:
AK: Conception, interpretation, and drafting of the manuscript.
MMG, IAM: Analysis and revision.
SHN, RS: Interpretation and revision.
SAHG: Conception and drafting of the manuscript.
All authors approved the final version of the manuscript to be published.

REFERENCES

1. CDC. Preventing leishmaniasis [Internet]. Leishmaniasis. 2024. Available from: https://www.cdc.gov/leishmaniasis/ prevention/index.html.
2. Knight CA, Harris DR, Alshammari SO, Gugssa A, Young T, Lee CM. Leishmaniasis: Recent epidemiological studies in the Middle East. Front Microbiol 2023; 13:1052478. doi: 10.3389/fmicb.2022.1052478.
3. Ayene YY, Mohebali M, Hajjaran H, Akhoundi B, Shojaee S, Rahimi-Foroushani A, et al. A comparative study of nested-PCR and direct agglutination test (DAT) for the detection of Leishmania infantum infection in symptomatic and asymptomatic domestic dogs. BMC Res Notes 2021; 14(1):270. doi: 10.1186/s13104-021-05654-0.
4. Arnau A, Abras A, Ballart C, Fernandez-Arevalo A, Torrico MC, Tebar S, et al. Evaluation of the diagnostic sensitivity of the VIASURE Leishmania real-time PCR detection kit prototype for the diagnosis of cutaneous and visceral leishmaniasis. Transbound Emerg Dis 2023; 2023:1-8. doi: 10.1155/2023/1172087.
5. Mathison BA, Bradley BT. Review of the clinical presentation, pathology, diagnosis, and treatment of leishmaniasis. Lab Med 2023; 54(4):363-71. doi: 10.1093/ labmed/lmac134.
6. Pagniez J, Petitdidier E, Parra-Zuleta O, Pissarra J, Bras-Goncalves R. A systematic review of peptide-based serological tests for the diagnosis of leishmaniasis. Parasite 2023; 30:10. doi: 10.1051/parasite/2023011.
7. Hussain M, Munir S, Khan TA, Khan A, Ayaz S, Jamal MA, et al. Epidemiology of cutaneous leishmaniasis outbreak, Waziristan, Pakistan. Emerg Infect Dis 2018; 24(1):159-61. doi: 10.3201/eid2401.170358.
8. Ahmad S, Obaid MK, Taimur M, Shaheen H, Khan SN, Niaz S, et al. Knowledge, attitude, and practices towards cutaneous leishmaniasis in referral cases with cutaneous lesions: A cross-sectional survey in remote districts of southern Khyber Pakhtunkhwa, Pakistan. PLoS One 2022; 17(5):e0268801. doi: 10.1371/journal.pone.0268801.
9. Naseer U, Muzafar S, Mian Sayed K. A brief review on infes-tation of cutaneous leishmaniasis in Pakistan. Biomed J Sci Tech Res 2020; 31(4):24405-8. doi: 10.26717/BJSTR. 2020.31.005141.
10. Gurel MS, Tekin B, Uzun S. Cutaneous leishmaniasis: A great imitator. Clin Dermatol 2020; 38(2):140-51. doi: 10. 1016/j.clindermatol.2019.10.008.
11. Roy M, Ceruti A, Kobialka RM, Roy S, Sarkar D, Wahed AAE, et al. Evaluation of recombinase polymerase amplification assay for monitoring parasite load in patients with kala-azar and post kala-azar dermal leishmaniasis. PLoS Negl Trop Dis 2023; 17(4):e0011231. doi: 10.1371/journal.pntd. 0011231.
12. Sousa AQ, Pompeu MM, Frutuoso MS, Lima JW, Tinel JM, Pearson RD. Press imprint smear: A rapid, simple, and cheap method for the diagnosis of cutaneous leishmaniasis caused by Leishmania (Viannia) braziliensis. Am J Trop Med Hyg 2014; 91(5):905-7. doi: 10.4269/ajtmh.14-0160.
13. Mesa LE, Manrique R, Muskus C, Robledo SM. Test accuracy of polymerase chain reaction methods against conventional diagnostic techniques for cutaneous leishmaniasis (CL) in patients with clinical or epidemiological suspicion of CL: Systematic review and meta-analysis. PLoS Negl Trop Dis 2020; 14(1):e0007981. doi: 10.1371/journal.pntd.0007981.
14. Buderer NM. Statistical methodology: I. Incorporating the prevalence of disease into the sample size calculation for sensitivity and specificity. Acad Emerg Med 1996; 3(9): 895-900. doi: 10.1111/j.1553-2712.1996.tb03538.x.
15. Bain BJ, Bates I, Laffan MA. Dacie and lewis practical haema-tology. ed. 12^th, London, UK: Elsevier Ltd.; 2017. Available from: https://www.sciencedirect.com/book/9780702066962/ dacie-and-lewis-practical-haematology.
16. Bacterial xpress nucleic acid extraction kit [Internet]. MERCK. Available from: https://www.merckmillipore.com/ INTL/en/product/Bacterial-Xpress-Nucleic-Acid-Extraction-Kit,MM_NF-3096.
17. Chang KP, Fish WR. In vitro cultivation of protozoan parasites. In: Jenson JB, Ed. ed. 1^st, London, UK: CRC Press; 1983.
18. Umemneku Chikere CM, Wilson KJ, Allen AJ, Vale L. Comparative diagnostic accuracy studies with an imperfect reference standard - A comparison of correction methods. BMC Med Res Methodol 2021; 21(1):67. doi: 10.1186/ s12874-021-01255-4.
19. Al-Jawabreh A, Schoenian G, Hamarsheh O, Presber W. Clinical diagnosis of cutaneous leishmaniasis: A comparison study between standardized graded direct microscopy and ITS1-PCR of Giemsa-stained smears. Acta Trop 2006; 99(1):55-61. doi: 10.1016/j.actatropica.2006.07.001.
20. Ullah Z, Samad F, Bano R, Arif S, Zamir S, Aziz N, et al. Characterizing cutaneous leishmaniasis in a conflict-affected region: A study from North Waziristan, Pakistan. Turk J Med Sci 2023; 53(6):1767-75. doi: 10.55730/1300- 0144.5746.
21. Nateghi Rostami M, Darzi F, Farahmand M, Aghaei M, Parvizi P. Performance of a universal PCR assay to identify different Leishmania species causative of old world cutaneous leishmaniasis. Parasit Vectors 2020; 13(1):431. doi: 10.1186/s13071-020-04261-5.
22. Ali MA, Khamesipour A, Rahi AA, Mohebali M, Akhavan A, Firooz A, et al. Epidemiological study of cutaneous leishmaniasis in some Iraqi provinces. J Men's Health 2018; 14(4):e18-e24. doi:10.22374/1875-6859.14.4.4.