Special Issue:

Advancements in Animal Health and Production in Low and Middle-Income Countries

Detection of Avian Malaria (Plasmodium spp.) in Domestic Local Breed Chickens (Gallus gallus domesticus) Using Real Time PCR

Rana Mohammed Ibrahim

Department of Microbiology, College of Veterinary Medicine, University of Baghdad.

Received | July 12, 2025; Accepted | August 18, 2025; Published | September 03, 2025

*Correspondence | Rana Mohammed Ibrahim, Department of Microbiology, College of Veterinary Medicine, University of Baghdad; Email: [email protected]

Citation | Ibrahim RM (2025). Detection of avian malaria (Plasmodium spp.) in domestic local breed chickens (Gallus gallus domesticus) using real time PCR. J. Anim. Health Prod. 13(s1): 293-299.

DOI | https://dx.doi.org/10.17582/journal.jahp/2025/13.s1.293.299

ISSN (Online) | 2308-2801

Copyright: 2025 by the authors. Licensee ResearchersLinks Ltd, England, UK.

This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/).

Introduction

Haemosporidian protozoan parasites that infect mammals, birds, reptiles, and amphibians (Valkiünas, 2005). The greater majority of avian Plasmodium spp. description was reported in birds that live in tropical and subtropical countries and have been adapted for transmission to become cosmopolitan (Valkiünas and Iezhova, 2018). Plasmodium species found in poultry such as P. durae, P. gallinaceum, P. relictum, and P. juxtanucleare (Santiago-Alarcon et al., 2020; Zajac et al., 2021). They are causing many diseases, such as avian malaria (Beadell et al., 2004; Hellgren et al., 2004), which may lead to severe mortalities in domestic, wild, and zoo birds (Ferrell et al., 2007). Avian Plasmodium spp. has a much broader host range and frequently occurs in several avian families (Krizanauskiene et al., 2006). Of the 13 blood-feeding families, four (Culicidae, Simuliidae, Ceratopogonidae, and Hippoboscidae) are known to be involved as vectors of avian haemosporidian parasites, and an overview of their morphology, biology, importance, taxonomy, and diversity is presented (Ibáñez-Bernal et al., 2020). The life cycle of a parasite requires two hosts, typically a bird and a female mosquito, which most often belong to the genus Culex (or Culicoides midges) (Atkinson and van Riper III, 1991; Valkiünas, 2005). It proliferates as clones, similar to haploid in bird hosts, and through asexual reproduction in vectors (Desser and Bennett, 1993). The clinical symptoms of the disease differ according to species. The mild symptoms, such as paleness of the comb, wattle, and lethargy, and the infection of P. juxtanucleare can be moderate, mild, or asymptomatic (Kissinger et al., 2002; Vaisusuk et al., 2022) and/or may exhibit lethargy, white diarrhea, and convulsions (Tattiyapong et al., 2016), and in older chickens, it is mild paleness of the comb, wattle, and lethargy (Dhamayanti et al., 2023; Mohsen et al., 2024).

A different traditional diagnostic methods are used for detection of Plasmodium spp. which is the only undergo schizogony in circulating blood cells, that presence inside the erythrocytes and the both schizonts and gametocytes help in diagnoses of the parasite infections (Atkinson and van Riper III, 1991), such as blood smears or by using recent developmental methods (molecular technologies) as a more reliable (Fallon et al., 2003). Over the last two decades, molecular methods for studying mitochondrial DNA sequence variation have become an integral part of avian haemosporidian research. These methods have primarily been used for identification of the parasites and tentative phylogenetic reconstructions, allowing to compare data across the globe (Bensch and Hellgren, 2020), for an indication of the high diversity (Ricklefs et al., 2007; Kim and Tsuda, 2010) and identify the species of the parasites by sequencing of the amplified DNA (Bensch et al., 2009). Most investigations into avian malaria have been primarily based on material collected from naturally infected birds, and important data on ecology, distribution, molecular biology, diversity, and phylogeny have been accumulated since the end of the 20th century (Beadell et al., 2006; Hellgren et al., 2007). The model for malaria research is represented in chickens due to the severity of the disease, its pathology, and the unique model it provides for understanding the evolution of the parasite and its ecology in both field and laboratory settings (Macchi et al., 2010).

To the best of our knowledge, no previous molecular real time PCR analysis has been conducted to detect avian malaria (Plasmodium spp.) in domestic local breed chickens (Gallus gallus domesticus). This study not only offer a molecular detection system but also highlight the need of continued monitoring of the infection in birds.

Materials and Methods

Blood samples

Forty-five local breed chickens (Gallus gallus domesticus) were purchased from different local markets in Baghdad city during the period from 1/10/2021 to 1/5/2022. About 1-2 ml from jugular vein was collected in EDTA (Ethylene diamine tetra acetic acid) tubes (Al-Daraji et al., 2008) and they were kept in deep freezer at -20 oC until analysis by real time polymerase chain reaction in the Laboratory of Microbiology Department, College of Veterinary Medicine, University of Baghdad.

DNA extraction from blood

Approximately 25 µL of blood was transferred into a 1.5 mL tube. The freezing-thawing was used for DNA extraction, added 200 µl buffer CL, 20 μl Proteinase K, and 5 μl RNase A solution into the sample tube and mixed vigorously, incubated the lysate at 56℃ using a preheated heat block for 10 ~ 30 min., added 200 μl of Buffer BL into the upper sample tube and mixed thoroughly, then incubated at 70℃ for 5 min., centrifuged at 13000 rpm for 5 min, and carefully transferred 350 ~ 400 μl of the supernatant into a new 1.5 ml tube and centrifuged. Added 200 μL of absolute ethanol to the lysate in the 1.5 mL tube, then centrifuged. The mixture was then applied to the spin column and centrifuged at 13,000 rpm for 1 min. Discard the filtrate and place the spin column in a new 2 ml collection tube. Added 700 μl of buffer WB to the Spin column and centrifuged for 1 min.at 13000 rpm. Placed the column into a new 2.0 mL collection tube and then centrifuged for 1 minute. Discarded the flow-through and collection tube altogether. Placed the spin column into a new 1.5 ml tube and added 30-100 μl of Buffer CE directly onto the membrane. Incubated for 1 min at room temperature and then centrifuged for 1 min at 13000 rpm to elute.

The primers preparation

The lyophilized, designed primers (AMA 1 gene) with their IDs were dissolved in free ddH2O to give a final concentration of 100 pmol/µL (stock solution) and stored at -20 °C. Prepared 10 pmol/µl concentration as working primer suspended as shown in the table below:

and the optimum conditions for gene detection were illustrated below:-

No.	Phase	Tm (ᵒC)	Time	No. of cycle
1-	Initial Denaturation	95ᵒC	5 min	1 cycle
2-	Denaturation -2	95ᵒC	45 Sec	35 cycle
3-	Annealing	52ᵒC	45 Sec
4-	Extension-1	72ᵒC	1min
5-	Extension -2	72ᵒC	5 min.	1 cycle

 

Primer	Sequence	Primer sequence	Tm	GC %	Size of product (bp)	Reference
(AMA-1) gene	F	5'- GTCTTTGAATTAAGTGCTTCGGA - 3'	57.39	39.13	200 bp	Primer design (Id: LT969560.1)
	R	5'- TGTGTTACGGTGTTATAATTTCCCC - 3’	59.35	40.00

 

Gel electrophoresis

Agarose (1.5%) was used to determine the DNA according to Sambrook et al. (1989).

Real time PCR

The KAPA SYBR FAST qPCR Kit is supplied as a 2X master mix with integrated antibody-mediated hot start, SYBR Green I fluorescent dye, MgCl2, dNTPs, and stabilizers. The optimum conditions of the RT-PCR cycling program for gene detection are as follows:

No.	Phase	Tm (ᵒC)	Time	No. of cycles
1-	Enzyme activation	95	5 min.	Hold
2-	Denaturation -2	95	0.2 Sec.	40
3-	Annealing	56	0.2 Sec.
4-	Extension-1	72	0.2sec.
5-	Final Extension -2	72	2 min.	1
6	Melting curve	90	0.15 sec.	1

 

Statistical analysis

Statistical analysis was performed according to Al-Mohammed et al. (1986) using the Chi-square test to detect significant differences in the effect of specific epidemiological parameters (sex and location) of the study at a significance level of P ≤ 0.05 or 0.01.

Results

The total infection rate of Plasmodium spp. in local breed chickens (Gallus gallus domesticus) was 15.55% (7/45) in Baghdad city (Figures 1 and 2).

 

The infection rate among males was 15% (3/20), while among females it was 16% (4/25), with a significant difference (P ≤ 0.01) (Table 1).

According to the area, a highly significant (P ≤ 0.01) infection rate was observed in the Al-Karkh district market at 18.18% (4/22), compared to the Al-Rusafa district market at 13.03% (3/23) (Table 2).

 

Table 1: The infection rate of Plasmodium spp. in local breed chickens (Gallus gallus domesticus) at Baghdad city.

Sex	No. of chickens examined	Positive (%)
Males	20	3(15)
Females	25	4(16)
Total	45	7(15.55)
χ2	10.23 ** P ≤ 0.01

 

Table 2: The infection rate of Plasmodium spp. in local breed chickens (Gallus gallus domesticus) at Baghdad city according to the district markets.

Districts markets	No. of chickens examined	Positive (%)
Al-Rusafa	23	3(13.04)
Al-Karkh	22	4(18.18)
Total	45	7(15.55)
χ2	11.20 ** P≤0.01

 

Discussion

Blood parasite infections in poultry, such as those caused by Plasmodium, pose a serious threat to the poultry industry due to their potential to cause significant economic losses (Dhamayanti et al., 2023). Morphological identification using microscopic examination of blood films remains an important tool for malaria diagnostics. It is particularly valuable when applied in conjunction with PCR as a diagnostic tool (Valkiünas and Iezhova, 2018). PCR is used as a molecular marker, which is sensitive for distinguishing different parasite species and their lineages, and is essential for identifying cryptic Plasmodium species (Palinauskas et al., 2015).

Microscopic examination revealed that 17/105 positives (16.19%) were positive for blood parasite infection. Trophozoites, erythrocytic meronts, and microgametocytes of Plasmodium were found in blood samples. Based on morphological examination, the species found in the samples was closely related to P. juxtanucleare. Polymerase chain reaction examination revealed that 21 out of 60 samples were positive for Plasmodium (35%). The Plasmodium species identified from the sequenced samples were proven to be P. juxtanucleare, which is endemic to Thailand, and it is closely related to the samples (99.64%–100%), with a genetic distance of 0%–1%.

Additionally, age, population, and cage type were not found to be significantly associated with Plasmodium infection (Dhamayanti et al., 2023). The adaptation of chicken immunity to P. juxtanucleare infection may influence the severity of infection (Vaisusuk et al., 2022). Chicken immunity can also be affected by cage type; chickens in battery cages showed lower immunity due to chronic stress compared to those in enriched cages. On the other hand, chickens kept in enriched cages and those that are free-range tend to have a higher chance of contracting the disease (Bhanja and Bhadauria, 2018; Hofmann et al., 2020). In chickens that survive Plasmodium infection, they still experience low parasiteemia even when the parasiteemia level does not increase (Paulman et al., 2005). Samples that underwent PCR showed more positive results than those that underwent microscopic examination, and polymerase chain reaction has higher sensitivity than microscopic examination (Dhamayanti et al., 2023).

The thickness of the band on the agarose gel for each sample indicated the level of parasitemia. The lowest parasitaemia level was 0.21%, represented by the thinnest band. In contrast, the highest parasiteamia level among the positive samples was 0.64%, represented by the thickest band, and the thickness of the band on the agarose gel for each sample indicated the parasiteamia level (Dhamayanti et al., 2023). Bensch et al. (2009) demonstrated a correlation between the thickness of the band on an agarose gel and the rate of erythrocyte infection, with the thinnest band observed at 0.04% and the thickest at 1.93%.

The host-specificity of avian malaria parasites is diverse: some parasites can infect hosts from multiple families or even multiple orders; others are restricted to a single avian family or even species (Ricklefs and Fallon, 2002; Fallon et al., 2005; Loiseau et al., 2012).

Of the 55 samples tested using PCR analysis, 39 (71%) were positive for Plasmodium spp., and 28/40 (62%) blood smears were positive for Plasmodium spp. This is the first report from New Zealand in which specific Plasmodium spp. mixed infections have been found in introduced birds. Co-infections with several cosmopolitan Plasmodium lineages were identified, as well as the first report in New Zealand of an exotic avian Plasmodium sp. lineage, in Australian magpies. Whilst the role of introduced birds in maintaining and spreading pathogenic avian malaria in New Zealand is unclear, there is a potential infection risk to native birds, especially where distributions overlap (Schoener et al., 2019).

Ibrahim and Al-Rubaie (2020) were reported that the prevalence of avian malaria (Plasmodium gallinaceum) was 18% (18/100), which divided closely between males (20.00%) and females 16.00%, with a moderate spread in the local domesticated breed chickens (Gallus gallus domesticus) in Baghdad city. Shadan (2013) reported that the most prevalent Haemoparasite was Plasmodium spp., with an infection rate of 52.63% (70/133), while Flayyih (2014) recorded an infection rate of 9.86% in Turkey. This study found no relationship between the age of the chickens, the type of cage, and the population, which may be due to the insufficient sample size. Based on microscopic and PCR examinations, the Plasmodium species found in layer chickens from the three districts of Yogyakarta, Indonesia, was identified as P. juxtanucleare, which showed a low genetic distance from P. juxtanucleare from Thailand and Japan (0%–1%). There was no correlation between Plasmodium infection and age, type of cage, or population of layer chickens, but this may be due to the insufficient sample size (Dhamayanti et al., 2023). Avian malaria and related haemosporidian parasites are globally distributed, with host diversity and environmental factors in tropical regions favoring the prevalence and diversity of this group of parasites (Santiago-Alarcon and Marzal, 2020). A combination of microscopic, PCR-based, and serological diagnostic methodologies provides better estimates of the proper distribution and other aspects of the biology of haemosporidians, particularly in studies on virulence, prevalence, and biodiversity (Valkiünas and Atkinson, 2020). These parasites were monitored by microscopic analysis of thin blood smears stained with Giemsa stain (Prunk-Nern et al., 2014). However, it is not a reliable method when performed by non-experts due to a lack of specialized training and expertise, as it requires specific training and considerable expertise (Chen et al., 2013; Razzak and Al-Haqban, 2015). Despite avian malaria having been detected by microscopic examination of blood smears (Greiner et al., 1975; McClure et al., 1978; Peirce, 1981; Atkinson and Van Riper III, 1991). Differences may be related to the diversity of the vectors (Culex, Aedes, and Simulium, which are abundant in rivers/or to the specimens that were collected (Cohen, 1977). Haemoparasite infections mainly depend on invertebrate vectors, usually sucking insects, to infect their avian host (Valkiünas, 2005). In Plasmodium parasites that infect multiple host species through host-switching events, such as avian malaria species, these polymorphisms may contribute to a broad range of host specificity (Iyer et al., 2007). The effects influence the host’s habitat choice, and bird species that are unable to develop resistance to these parasites may have increased vectors and abundance, which in turn increase the chances of infection (Atkinson et al., 1995; 2000; Woodworth et al., 2005). The total infection rate, 28.33% (51/180), was recorded in Plasmodium spp. in chickens (Gallus gallus domesticus) using Giemsa-stained blood smears, with a higher infection rate in females than in males, but without significant differences (Ibrahim and A-Rubaie, 2020). Zamaura-Vilchis et al. (2012) found that 1.74% Plasmodium is found in birds. Fernandez-Davila and Phalen (2013) found 10.9% Plasmodium in birds. The differences between the results of the current study and the previous studies in the infection rates may be due to the diagnostic methods or to parasite biological habitat as heteroxenous parasite and need more than one obligatory host in its life cycle, which are dipteran insects transmit the infective stages between birds hosts (Atkinson and Van Riper III, 1991; Valkiünas, 2005; Angrisano et al., 2012).

ACKNOWLEDGEMENT

The authors express their gratitude to the College of Veterinary Medicine, Department of Microbiology, University of Baghdad for their invaluable assistance in successfully conducting the study. We acknowledge the support of time and facilities from for this study

NOVELTY STATEMENT

This study introduces a novel approach to detecting Plasmodium spp. infections in domestic local breed chickens (Gallus gallus domesticus) in Baghdad, utilizing Real-Time PCR for enhanced sensitivity and specificity. By examining 45 birds from various regional markets, the research not only addresses a significant gap in understanding avian malaria prevalence in this region but also provides crucial insights into the potential impact of such infections on poultry health and local agriculture. This work contributes to the broader field of veterinary parasitology and offers a foundation for future studies on disease management in poultry populations.

AUTHOR’S CONTRIBUTION

Rana Mohammed Ibrahim conducted the study design, data collection, analysis, and manuscript preparation. The author is solely responsible for all aspects of the work.

Generative AI or AI-assisted Technology Statement

The author(s) declare that no Genrative AI was used in the creation of this manuscript.

Conflict of interest

The authors has declared no conflict of interest.

References

Al-Daraji HJ, Al-Hayani WK, Al-Hassani AS (2008). Avian hematology. College of Agriculture, University of Baghdad. pp. 2–10.

Al-Mohammed NT, Al-Rawi KM, Younis MA, Al-Morani WK (1986). Principles of statistics. Press Baghdad University.

Angrisano F, Tan YH, Sturm A, McFadden GI, Baum J (2012). Malaria parasite colonization of the mosquito midgut Placing the Plasmodium ookinete center stage. Int. J. Parasitol., 42: 519–527. https://doi.org/10.1016/j.ijpara.2012.02.004

Atkinson CT, van Riper III C (1991). Pathogenicity and epizootiology of avian hematozoa: Plasmodium, Leucocytozoon and Haemoproteus. In: (eds. J. Loye and M. Zuk). Bird-parasite interactions: Ecology, evolution, and behavior. Oxford University Press, New York: pp. 19–48. https://doi.org/10.1093/oso/9780198577386.003.0002

Atkinson CT, Woods KL, Dusek RJ, Sileo LS, Iko WM (1995). Wildlife disease and conservation in Hawaii: Pathogenicity of avian malaria Plasmodium relictum in experimentally infected Iiwi Vestiaria coccinea. Parasitology, 111(S1): S59–S69. https://doi.org/10.1017/S003118200007582X

Atkinson, C.T., R. J. Dusek, K.L.Woods, W.M.IKO (2000).pathogenicity of avian malaria in experimentally infected Hawaii Amakihi.J.Wildl.Dis.,36 :197_204

Beadell JS, Gering E, Austin J, Dumbacher JP, Peirce MA, Pratt TK, Atkinson CT, Fleischer RC (2004). Prevalence and differential host-specificity of two avian blood parasite genera in the Australo-Papuan region. Mol. Ecol., 13: 3829–3844. https://doi.org/10.1111/j.1365-294X.2004.02363.x

Beadell JS, Ishtiaq F, Covas R, Melo M, Warren BH, Atkinson CT, Bensch S, Graves GR, Jhala YV, Peirce MA, Rahmani AR, Fonseca DM, Fleischer RC (2006). Global phylogeographic limits of Hawaii’s avian malaria. Proc. Biol. Sci. R. Soc., 273: 2935–2944. https://doi.org/10.1098/rspb.2006.3671

Bensch S, Hellgren O (2020). The use of molecular methods in studies of avian haemosporidians. In: Avian malaria and related parasites in the tropics: Ecology, evolution and systematics. 1st ed. Springer International Publishing, Cham. pp. 113–135. https://doi.org/10.1007/978-3-030-51633-8_4

Bensch S, Hellgren O, Pérez-Tris J (2009). MalAvi: A public database of malaria parasites and related haemosporidians in avian hosts based on mitochondrial cytochrome b lineages. Mol. Ecol. Resour., 9: 1353–1358. https://doi.org/10.1111/j.1755-0998.2009.02692.x

Bhanja S, Bhadauria P (2018). Behavior and welfare concepts in laying hens and their association with housing systems. Int. J. Poult. Sci., 53(1): 1–10. https://doi.org/10.5958/0974-8180.2018.00009.0

Chen T, Zhang Y, Wang C, Zhenshen Q, Wang F, Syeda-Mahmood TS (2013). Complex local phase based subjective surfaces (CLAPSS) and its application to DIC red blood cell image segmentation. Neurocomputing, 99: 98–110. https://doi.org/10.1016/j.neucom.2012.06.015

Cohen S (1977). Mechanisms of malaria immunity. Trans. R. Soc. Trop. Med. Hyg., 71: 283–286. https://doi.org/10.1016/0035-9203(77)90100-6

Desser SS, Bennett GF (1993). The genera Leucocytozoon, Haemoproteus, and Hepatocystis. In: Kreier JP (ed). Parasitic Protozoa. Vol. 7, 2nd ed. Academic Press, San Diego, CA: 273–310. https://doi.org/10.1016/B978-0-12-426014-6.50010-7

Dhamayanti E, Priyowidodo D, Nurcahyo W, Firdausy LW (2023). Morphological and molecular characteristics of Plasmodium juxtanucleare in layer chicken from three districts of Yogyakarta, Indonesia. Vet. World, 16(8): 1576–1583. https://doi.org/10.14202/vetworld.2023.1576-1583

Fallon SM, Bermingham E, Ricklefs RE (2005). Host specialization and geographic localization of avian malaria parasites: A regional analysis in the Lesser Antilles. Am. Nat., 165: 466–480. https://doi.org/10.1086/428430

Fallon SM, Ricklefs RE, Swanson BL, Bermingham E (2003). Detecting avian malaria: An improved polymerase chain reaction diagnosis. J. Parasitol., 89(5): 1044–1047. https://doi.org/10.1645/GE-3157

Fernandez-Davila,M.A., D. phalen (2013).Haemosporidia and Australian wild birds .awhn@ zoonsw.gov.au: 1_9.

Ferrell ST, Snowden K, Marlar AB, Garner M, Lung NP (2007). Fatal hemoprotozoal infections in multiple avian species in a zoological park. J. Zoo. Wildl. Med., 38(2): 309–316. https://doi.org/10.1638/1042-7260(2007)038[0309:FHIIMA]2.0.CO;2

Flayyih MM (2014). Isolation and identification of some ectoparasites, haemoprotozoa, Histomonas sp in turkey (Meleagris gallopavo) in Naseria city. MSc thesis, College of Veterinary Medicine, University of Baghdad: 76p.

Greiner EC, Bennett GF, White EM, Coombs RF (1975). Distribution of the avian hematozoa of North America. Can. J. Zool., 53: 1762–1787. https://doi.org/10.1139/z75-211

Hellgren O, Krizanauskiene A, Valkiunas G, Bensch S (2007). Diversity and phylogeny of mitochondrial cytochrome B lineages from six morphospecies of avian Haemoproteus (Haemosporida, Haemoproteidae). J. Parasitol., 93(4): 889–896. https://doi.org/10.1645/GE-1051R1.1

Hellgren O, Waldenstrom J, Bensch S (2004). A new PCR assay for simultaneous studies of Leucocytozoon, Plasmodium, and Haemoproteus from avian blood. J. Parasitol., 90(4): 797–802. https://doi.org/10.1645/GE-184R1

Hofmann T, Schmucker SS, Bessei W, Grashorn M, Stefanski V (2020). Impact of housing environment on the immune system in chickens: A review. Animals, 10(7): 1138. https://doi.org/10.3390/ani10071138

Ibanez-Bernal S, Rivera-Garcia KD, Abella-Medrano CA (2020). Introduction to the taxonomy and general biology of Diptera (Insecta) involved in the transmission of avian Haemosporida. In: Avian malaria and related parasites in the tropics: Ecology, evolution and systematics. 1st ed. Springer International Publishing, Cham. pp. 137-184. https://doi.org/10.1007/978-3-030-51633-8_5

Ibrahim RM, Al-Rubaie HM (2020). Molecular detection of avian malaria (Plasmodium gallinaceum) in local domesticated breed chickens (Gallus gallus domesticus) in Baghdad. Iraqi J. Vet. Med., 44(E0): 75-79. https://doi.org/10.30539/ijvm.v44i(E0).1025

Ibrahim RM, Al-Rubaie HM (2020). Prevalence of some haemosporidians in domesticated chickens in Baghdad city. Plant Arch., 20(Suppl 1): 3444-3448.

Iyer J, Gruner AC, Renia L, Snounou G, Preiser PR (2007). Invasion of host cells by malaria parasites: A tale of two protein families. Mol. Microbiol., 65: 231–249. https://doi.org/10.1111/j.1365-2958.2007.05791.x

Kim KS, Tsuda Y (2010). Seasonal changes in feeding pattern of Culex pipiens pallens govern the transmission dynamics of multiple lineages of avian malaria parasites in Japanese wild bird community. Mol. Ecol., 19(24): 5545-5554. https://doi.org/10.1111/j.1365-294X.2010.04897.x

Kissinger JC, Souza PC, Soares CO, Paul R, Wahl AM, Rathore D, McCutchan TF, Krettli AU (2002). Molecular phylogenetic analysis of the avian malarial parasite Plasmodium (Novyella) juxtanucleare. J. Parasitol., 88(4): 769–773. https://doi.org/10.1645/0022-3395(2002)088[0769:MPAOTA]2.0.CO;2

Krizanauskiene A, Hellgren O, Kosarev V, Sokolov L, Bensch S, Valkiunas G (2006). Variation in host specificity between species of avian hemosporidian parasites: Evidence from parasite morphology and cytochrome b gene sequences. J. Parasitol., 92: 1319-1324. https://doi.org/10.1645/GE-873R.1

Loiseau C, Harrigan RJ, Robert A, Bowie RCK, Thomassen HA, Smith TB, Sehgal RNM (2012). Host and habitat specialization of avian malaria in Africa. Mol. Ecol., 21: 431–441. https://doi.org/10.1111/j.1365-294X.2011.05341.x

Macchi BM, Quaresma JA, Herculano AM, Crespo-Lopez ME, DaMatta RA, Nascimento JL (2010). Pathogenic action of Plasmodium gallinaceum in chickens: Brain histology and nitric oxide production by blood monocyte-derived macrophages. Vet. Parasitol., 2: 16-22. https://doi.org/10.1016/j.vetpar.2010.04.032

McClure HE, Poonswad P, Greiner EC, Laird M (1978). Haematozoa in the birds of Eastern and Southern Asia. Memorial University of Newfoundland, St. John’s, Newfoundland, Canada.

Mohsen AK, Jassim SE, Alaridhi TT, Al-Erjan AM, Lahhob QR, Mudhafar M (2024). Revolutionizing infectious disease management in animals through advanced phage therapy: A comprehensive strategy for the eradication of multidrug-resistant bacterial pathogens. J. Anim. Health Prod., 12(s1): 157-167. https://doi.org/10.17582/journal.jahp/2024/12.s1.157.167

Palinauskas V, Ziegtye R, Ilgunas M, Iezhova TA, Bernotiene R, Bolshakov C, Valkiunas G (2015). Description of the first cryptic avian malaria parasite Plasmodium homocircumflexum n. sp., with experimental data on its virulence and development in avian hosts and mosquitoes. Int. J. Parasitol., 45(1): 51–62. https://doi.org/10.1016/j.ijpara.2014.08.012

Paulman A, Mcallister MM (2005). Plasmodium gallinaceum: Clinical progression, recovery, and resistance to disease in chickens infected via mosquito bite. Am. J. Trop. Med. Hyg., 73(6): 1104–1107. https://doi.org/10.4269/ajtmh.2005.73.1104

Peirce MA (1981). Distribution and host-parasite checklist of the haematozoa of birds in Western Europe. J. Nat. Hist., 15: 419-458. https://doi.org/10.1080/00222938100770321

Prunk-Nern M, Pattaradilokrat S, Chumpolbanchorn K, Pimnon S, Narkpinit S, Harnyu-ttanakorn P, Buddhirakkul P, Saiwichai T (2014). Effects of artesunate treatment on Plasmodium gallinaceum transmission in the vectors Aedes aegypti and Culex quinquefasciatus. Vet. Parasitol., 207(1-2): 161-165. https://doi.org/10.1016/j.vetpar.2014.10.032

Razzak MI, AlHaqban IB (2015). Automatic detection of malarial parasite using microscopic blood images. J. Med. Imag. Hlth. Inf., 5: 1-8. https://doi.org/10.1166/jmihi.2015.1417

Ricklefs RE, Fallon SM (2002). Diversification and host switching in avian malaria parasites. Proc. R. Soc. Lond. B, 269: 885–892. https://doi.org/10.1098/rspb.2001.1940

Ricklefs RE, Swanson BL, Fallon SM, Martinez-Abrain A, Scheuerlein A, Gray J, Latta SC (2007). Community relationships of avian malaria parasites in Southern Missouri. Ecol. Monogr., 75: 543-559. https://doi.org/10.1890/04-1820

Sambrook J, Fritschi EF, Maniatis T (1989). Molecular cloning: A laboratory manual 2nd ed. Cold Spring Harbor Laboratory Press, New York. University of Texas, South Western Medical Center, USA

Santiago-Alarcon D, Marzal A (2020). Research on avian haemosporidian parasites in the tropics before the year 2000. In: Avian malaria and related parasites in the tropics: Ecology, evolution and systematics. 1st ed. Springer International Publishing, Cham. pp. 1-44. https://doi.org/10.1007/978-3-030-51633-8_1

Schoener ER, Tompkins DM, Parker KA, Howe L, Castro I (2019). Presence and diversity of mixed avian Plasmodium spp. infections in introduced birds whose distribution overlapped with threatened New Zealand endemic birds. N. Z. Vet. J., 68(2): 1-14. https://doi.org/10.1080/00480169.2019.1680326

Shadan HA. (2013). Prevalence of blood parasites in local chickens in Qaradagh District,Sulaimani ,Iraq.Iraqi J. Vet. Med. 2013; 37(1):17_21.

Silveria P, Marin SYG, Moreira PA, Tocantins BB, Lacorte G, Paixao TA, Martins NRS, Braga EM (2013). Interactions of Plasmodium juxtanucleare and chicken anemia virus: Establishing a model. Parasitology, 140(14): 1777–1778. https://doi.org/10.1017/S0031182013001170

Tattiyapong, M., Deemagarn, T. and Jiratanh, S. Molecular (2016). characterization of plasmodium juxtanucleare in Burmese red jungle fowls (Gallus gallus spadiceus) in Thailand J.orotozool. Res., 26: 1_10.

Vaisusuk K, Chatan W, Seerintra T, Piratae S (2022). High prevalence of Plasmodium infection in fighting cocks in Thailand determined with a molecular method. J. Vet. Res., 66(3): 373–379. https://doi.org/10.2478/jvetres-2022-0049

Valkiunas G (2005). Avian malaria parasites and other haemosporidia. CRC Press, London. Boca Raton, Florida. pp. 936. https://doi.org/10.1201/9780203643792

Valkiunas G, Atkinson CT (2020). Introduction to life cycles, taxonomy, distribution, and basic research techniques. In: Avian malaria and related parasites in the tropics: Ecology, evolution and systematics. 1st ed. Springer International Publishing, Cham. pp. 45-80. https://doi.org/10.1007/978-3-030-51633-8_2

Valkiunas G, Iezhova TA (2018). Keys to the avian malaria parasites. Malar. J., 17(212): 1-24. https://doi.org/10.1186/s12936-018-2359-5

Woodworth BL, Atkinson CT, LaPointe DA, Hart PJ, Spiegel CS, Tweed EJ, Henneman C, LeBrun J, Denette T, DeMots R, Kozar KL, Triglia D, Lease D, Gregor A, Smith T, Duffy D (2005). Host population persistence in the face of introduced vector-borne diseases: Hawaii amakihi and avian malaria. Proc. Natl. Acad. Sci., 102(5): 1531-1536. https://doi.org/10.1073/pnas.0409454102

Zajac AM, Conboy GA, Little SE, Reichard MV (2021). Veterinary clinical parasitology. 9th ed. New Jersey: John Wiley and Sons, Inc.

Zamora-Vilchis I, Williams SE, Johnson CN (2012). Environmental temperature affects prevalence of blood parasites of birds on an elevation gradient: Implications for disease in a warming climate. PLoS One, 7: e39208. https://doi.org/10.1371/journal.pone.0039208