Next Article in Journal
Impact of the Gut Microbiota–Metabolite Axis on Intestinal Fatty Acid Absorption in Huainan Pigs
Previous Article in Journal
Single Mutation in iolT1 in ptsG-Deficient Corynebacterium glutamicum Enables Growth Boost in Xylose-Containing Media
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Microorganisms
Volume 13
Issue 7
10.3390/microorganisms13071607
microorganisms-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Occurrence and Quantification of Porcine Hemotrophic Mycoplasmas in Blood-Sucking Stomoxys calcitrans

by
Mareike Arendt
1,
Katharina Hoelzle
1,
Julia Stadler
2,
Mathias Ritzmann
2,
Julia Ade
1,
Ludwig E. Hoelzle
1,* and
Lukas Schwarz
3
1
Institute of Animal Science, Department of Livestock Infectiology and Environmental Hygiene, University of Hohenheim, 70593 Stuttgart, Germany
2
Clinic for Swine, Centre for Clinical Veterinary Medicine, Ludwig-Maximilians-Universität München, 85764 Oberschleissheim, Germany
3
Clinical Centre for Population Medicine in Fish, Pig and Poultry, Clinical Department for Farm Animals and Food System Science, University of Veterinary Medicine Vienna, 1210 Vienna, Austria
*
Author to whom correspondence should be addressed.
Microorganisms 2025, 13(7), 1607; https://doi.org/10.3390/microorganisms13071607
Submission received: 16 April 2025 / Revised: 4 July 2025 / Accepted: 5 July 2025 / Published: 8 July 2025
(This article belongs to the Section Veterinary Microbiology)
Review Reports Versions Notes

Abstract

Hemotrophic mycoplasmas (HMs) are cell wall-less, small and uncultivable pathogens, which can cause infections in pigs with no to severe clinical signs and can contribute to significant economic losses in the pig industry. In addition to the known mechanical transmission routes of HMs (e.g., via blood-contaminated instruments or lesions from ranking fights), transmission to pigs by arthropod vectors such as Stomoxys calcitrans is being discussed. To date, there is scant available data concerning the transmission of HMs by stable flies. The objective of this study is to gain more data concerning the occurrence of HMs in Stomoxys calcitrans. Therefore, quantitative real-time PCR was conducted on different stable fly samples (surface washings and whole flies). We found Mycoplasma (M.) suis in 5.2% of crushed flies and 4.2% of fly wash solutions, and M. parvum was detected in 5.2% of flies and 9.4% of fly wash solutions. ‘Candidatus (Ca.) M. haemosuis’ was not detected in any sample. The mean bacterial loads were 2.0 × 102 M. suis/fly, 9.3 × 102 M. suis/fly wash solution and, for M. parvum, 2.4 × 103 M. parvum/fly and 2.1 × 103 M. parvum/fly wash solution. This molecular occurrence of porcine HMs in blood-sucking flies and reasonable bacterial loads in the two- to three-digit range demonstrate that these flies serve as mechanical vectors in stables and are, therefore, of epidemiological importance.

1. Introduction

Hemotrophic mycoplasmas (HMs), which appear worldwide, cause significant economic losses to the pig industry [1,2,3]. Most HMs are host specific and can occur in a wide range of animals [4,5,6,7,8,9]. In pigs, three HM species, M. suis, M. parvum and the recently described ‘Ca. M. haemosuis’, are known so far. M. suis causes infectious anemia in pigs (IAP), which can manifest itself in various degrees of severity, depending on host susceptibility and virulence [10]. Pigs with acute infections show high fever, hemolytic anemia and hypoglycemia, whereas chronic infections vary from subclinical courses to anemia in neonates, growth retardation in feeder pigs and poor reproductive performance and dysgalactia in sows [3,10,11]. Contrary to infections with M. suis, M. parvum seems to be non-pathogenic, and infections are characterized by the absence of clinical signs or anemia [12]. ‘Ca. M. haemosuis’ has been detected in pigs with and without clinical signs in China, Korea, Germany and Thailand [13,14,15,16,17,18]. Clinical signs described include high fever, reduced feed intake, skin alterations, increased mortality and a decreased daily weight gain [17].
It has been shown that pigs can be infected with HMs through subcutaneous, intravenous, intramuscular, intraperitoneal and oral inoculation under experimental conditions [10,19,20]. Blood-independent infection routes, such as via excretions, seem to play a minor role under field conditions [21]. However, mechanical blood transmission through contaminated instruments, in utero transmission to newborns and lesions from ranking fights play an important role in infections [20,22,23]. In addition, arthropod vectors (e.g., stable flies, mosquitos, pig lice) are suspected to be involved in the transmission of porcine HMs. Though there is less data about the transmission of HMs by stable flies, in an infection experiment, healthy pigs were infected by stable flies (Stomoxys calcitrans) that had previously been blood-fed on M. suis infected pigs [23,24]. The obligate blood-sucking Stomoxys calcitrans, which can be found all over the world and is often present in pig farms, is also known to contribute to the mechanical transmission of other pathogens, such as Capripox virus, African swine fever virus, lumpy skin disease virus and Bacillus anthracis [25,26,27,28,29]. Due to defensive movements of the host after biting or due to other flies, the stable fly frequently changes hosts to continue its blood meal. Consequently, pathogens can be transferred, for instance, through their mouthparts or by regurgitation [30,31,32].
The aim of this study was to investigate the occurrence and bacterial loads of uncultivable HMs in stable flies (Stomoxys calcitrans) and to gain more knowledge about the vector potential of Stomoxys calcitrans for porcine HMs.

2. Material and Methods

2.1. Samples and Sample Preparation

Stable flies (Stomoxys calcitrans) from 20 Austrian piglet producing farms (stables 1–20) were available from a previous study [28]. Samples were collected mainly from the gestation area of farms with and without disease problems and subsequently screened for different bacteria and viruses including hemotrophic mycoplasma. Three flies from each pig farm were analyzed in this study.
Additional samples that originated from five Austrian pig farms with unknown HM status (stables 21–25), consisting of 10 blood samples and 6 flies per stable, were examined.
The Austrian farms were located in Lower Austria, Upper Austria, Styria and Vienna.
Furthermore, 6 stable flies from an HM-positive (M. suis, M. parvum and ‘Ca. M. haemosuis’) farm in Northern Germany, North Rhine-Westphalia, (stable 26) were tested for HMs.
Each stable fly sample was first washed in 400 µL Dulbecco’s Phosphate Buffered Saline (Biowest, Nuaillé, France). Then, the flies were individually washed in 70% ethanol, air dried and crushed with micropistills in a sterile 1.5 mL tube. DNA was isolated from the wash solutions and crushed flies using the GenEluteTM Bacterial Genomic DNA Kit (Sigma Aldrich, Darmstadt, Germany) according to the user’s manual, except for the following modification for crushed stable flies: Step 1 (centrifugation) was omitted, and the homogenized flies were mixed directly with 180 µL Lysis Solution T.
DNA from EDTA blood samples was extracted according to the manufacturers protocol using the Monarch® Genomic DNA Purification Kit (New England Biolabs, Frankfurt am Main, Germany) with the following preliminary steps to gain better HM DNA yield: a total of 200 µL EDTA blood was mixed with 200 µL lysis buffer (10 mM Tris-HCL pH 7.5, 5 mM MgCl2, 0.32 M saccharose, 1% (v/v) Triton X-100) and then centrifuged at 8000 rpm for 1 min. The supernatant was discarded. The pellet was resuspended in 400 µL lysis buffer and centrifuged as described above. This step was repeated once.

2.2. Quantitative Real-Time PCR

M. suis, M. parvum and ‘Ca. M. haemosuis’ were detected using the StepOneTM System (Applied Biosystems, Darmstadt, Germany). For detection of M. parvum, quantitative real-time PCR was performed as described elsewhere using primers MPaF and MPaR [33]. The samples were screened on M. suis by using the SYBR green assay and primers Msg1-Fw andMsg1-Rv targeting the MSG1 gene of M. suis as described by Hoelzle et al. [34]. ‘Ca. M. haemosuis’ was detected by quantitative real-time PCR using primers CMhsuisF and CMhsuisR to amplify a 177 bp fragment of the gap gene [35]. In each PCR, 2 µL DNA template was added to 18 µL PCR mixture containing 10 µL Fast SYBRTM Green Master Mix (Thermo Fisher Scientific, Darmstadt, Germany) and 8 µL primer mixture (containing 0.5 µM primer each for M. suis and ‘Ca. M. haemosuis’ and 0.4 µM primer each for M. parvum). Quantification was performed using plasmid DNA of M. suis, ‘Ca. M. haemosuis’ and M. parvum with standard dilutions 105, 104 and 103 genome equivalent (GE) per reaction in each qPCR run. By applying the Standard Curve Method of the StepOneTM software version 2.2 (Applied Biosystems), the obtained Ct values were then extrapolated into HM GE/reaction [33,34,35].
Further data analysis was performed using StepOneTM software version 2.2 (Applied Biosystems) and Microsoft® Excel, 2016.

2.3. Next Generation Sequencing

Two crushed fly samples, one that tested positive for M. suis and one that tested positive for M. parvum, were sent to Microsynth AG (Balgach, Switzerland) for next generation sequencing-based 16S rRNA analysis and subsequent data analysis including figure creation. Therefore, washing of flies and DNA isolation was performed as described above. Library preparation and sequencing, using the amplicon metagenomics approach, were then performed by Microsynth AG. By using the Illumina MiSeq system (Illumina, San Diego, CA, USA), they sequenced the hypervariable regions on the 16SrRNA gene, using universal primers 341F and 805R [36].

3. Results

3.1. Detection of HM Species by Quantitative Real-Time PCR

Mycoplasma suis was found in 4 out of 96 fly wash solutions (4.2%) and in 5 out of 96 fly samples (5.2%) originating from a total of six Austrian pig farms. Bacterial loads ranged from 7.7 × 102 to 1.1 × 103 M. suis in total positive wash solution samples (mean value 9.3 × 102 M. suis/washing solution sample) and 2.9 × 101 to 3.8 × 102 bacteria in positive fly samples (mean value 2.0 × 102 M. suis/fly sample). The results of M. suis-specific real-time PCRs are depicted in Table 1.
A total of 9 out of 96 wash solution samples (9.4%) tested positive for M. parvum, and 5 out of 96 fly samples (5.2%) were shown to be M. parvum positive. The M. parvum positive samples originated from a total of 12 stables. Bacterial loads ranged from 3.4 × 102 to 5.9 × 103 M. parvum/total washing solution sample (mean value 2.1 × 103 M. parvum/washing solution sample) and, in fly samples, from 1.2 × 103 to 3.9 × 103 M. parvum/fly sample (mean value 2.4 × 103 M. parvum/fly sample). The results of M. parvum-specific real-time PCRs are summarized in Table 2.
In 4 out of 26 stables (15.4%), both M. parvum and M. suis were detected in Stomoxys samples. In a total of 6 out of 26 stables (23.1%), M. suis positive flies were detected. M. parvum positive flies were found in 12/26 (46.2%). In HM positive stables, 28.9% (13/45) of the wash solution samples and 22.2% (10/45) of the fly samples tested positive.
‘Candidatus M. haemosuis’ was not detected in any of the Stomoxys samples.

3.2. Sequence Analyses

The results of the next generation sequencing-based 16S rRNA analysis (NGS) approach are summarized in Table 3. No HM specific sequences were identified by 16S rDNA NGS; only the presence of other bacteria, including Morganella morganii and Arthrobacter russicus, was detected.

4. Discussion

The epidemiology of porcine HM infections is rather unknown, and transmission by vectors is a recurring topic of discussion. The hematophagous stable fly Stomoxys calcitrans is often found in pig stables. Several studies have shown that Stomoxys calcitrans can transmit pathogens by mechanical transmission, carrying the pathogens on the body surface or via blood-feeding acts [31,37]. In the present study, we investigated Stomoxys calcitrans for the presence of the three porcine HM species M. suis, M. parvum and ‘Ca. M. haemosuis’ using species-specific real-time PCR assays. Because HMs are uncultivable, cultural methods cannot be considered for detection. The two HM species M. suis and M. parvum were present in the investigated flies, indicating that Stomoxys calcitrans could serve as a vector for these pathogens. Schwarz and co-workers examined pools of 10 flies out of some of the same stables (stables 1–20) for the presence of different pathogens including HMs. Using an HM universal PCR and subsequent sequence analyses, they identified 35% of the pig farms (7/20 farms) as positive for HMs [28]. The detected HM species also included M. suis and M. parvum. In this study, one farm with HM-positive stable flies had a history of IAP. However, at the time of sampling, the animals did not show any related clinical signs. In our study, HMs were found in 13 out of 26 stables (50%). In 4 of the 26 stables investigated, both M. parvum and M. suis were present in the flies. Differences in the detection rates between the present study and the study of Schwarz et al. [28] are most likely due to the investigation of single flies in our study instead of fly pools and the use of highly sensitive and species-specific real-time PCR assays. In our study, six fly samples from an HM-positive (M. suis, M. parvum and ’Ca. M. haemosuis’) farm in Northern Germany (stable 26) tested negative for HMs, indicating that large sample sizes of flies need to be analyzed to obtain information about the HM status of stables.
We analyzed surface wash solutions and whole flies in order to gain some insights into potential transmission modes. M. suis was present in 4.2% (4/96) of surface wash solutions and in 5.2% (5/96) of fly samples. M. parvum was present in 9.4% (9/96) of wash solutions and 5.2% (5/96) of whole Stomoxys calcitrans samples.
Positive wash solutions show that HMs are located on the surface of flies. There are essentially two possible ways on which these bacteria can get onto the surface of flies. Bacteria could either be transferred to the fly surface directly from infected pigs, or the fly surface could be contaminated via regurgitation of saliva or via fly feces. Furthermore, M. suis and M. parvum were detected in whole flies after thorough washing, indicating that the HMs are present in parts of the flies’ bodies. These results show that there are different possible modes of transmission. During the feeding act, Stomoxys can restart their blood meal on another host and hereby transmit pathogens from the surface of the fly into lesions of livestock [32]. Also, when injecting saliva prior to blood-sucking, they can inoculate with infected blood remaining on their mouthparts or regurgitated pathogens from their crop [31,38].
Interestingly, M. parvum was found more often in wash solutions than in the washed flies’ bodies, whereas for M. suis, the amounts of positive whole fly samples and wash solutions were similar. However, as the number of samples is very small, no final statement can be made as to whether there is a difference between the two species M. suis and M. parvum or whether the difference is a random result.
‘Candidatus M. haemosuis’ was not detected in any sample. Schwarz and co-workers were also not able to detect ‘Ca. M. haemosuis’ in the stable flies examined. However, the negative results regarding ‘Ca. M. haemosuis’ do not exclude the potential vector role of Stomoxys calcitrans, as ‘Ca. M. haemosuis’ is an emerging pathogen that has only been detected in domestic pigs from Asia (China, Republic of Korea and Thailand) and Germany so far [15,16,17,18,35]. In Austria, ‘Ca. M. haemosuis’ has not been confirmed yet, indicating a minor significance of this species in Austria so far. The overall lower prevalence of ‘Ca. M. haemosuis’ could be an explanation for the lack of detection of ‘Ca. M. haemosuis’ in the Stomoxys samples analyzed in the present study.
Arthropod contact as a potential transmission route for porcine HM is also supported by a previous study from Song and co-workers. The study proved that the prevalence of M. suis in summer and autumn was significantly higher than in other seasons. Thus, the air humidity in these seasons is more favorable for insects, and the study concluded that arthropod vectors might play a role in the transmission of M. suis [39]. In addition to the above-mentioned study by Schwarz and co-workers [28], only one previous study investigated the potential of Stomoxys in the experimental transmission of porcine HMs. Prullage and co-workers showed, in their infection study, that Stomoxys calcitrans fed on M. suis-infected pigs were able to infect splenectomized pigs when transferred immediately [24]. With longer intervals between feeding and contact with splenectomized pigs, the pigs could no longer be infected. Due to their lack of ability to maintain the pathogen, the study also concluded that Stomoxys calcitrans flies serve as mechanical vectors. In contrast, biological vectors can transmit pathogens after significant delay, as they are able to maintain the pathogen [24]. We could prove that HMs can be found in wash solutions of fly samples and crushed fly samples. Therefore, our results also suggest that Stomoxys calcitrans might serve as a mechanical vector for M. suis and M. parvum and transmit the bacteria either via surface contact or during the blood meal.
All other studies regarding the HM vector potential of Stomoxys calcitrans in livestock investigated bovine, ovine and buffalo HM species. One previous study detected the bovine HM species M. wenyonii in 41.41% of stable flies collected from a buffalo farm in Thailand [18]. Hornok and co-workers detected M. wenyonii and ‘Ca. M. haemobos’ DNA in Stomoxys calcitrans flies collected from a stable with HM-infected cattle [40]. One experimental study on ovine HMs was conducted by Overas in 1969, where Stomoxys spp. were used to successfully transfer M. ovis from infected to uninfected sheep [41]. However, in this study, maintenance and propagation of HMs in the vector were not included. Most studies investigating the potential of Stomoxys calcitrans as an HM vector for livestock are based on molecular detection methods due to HMs being uncultivable [42]. Although there was no evidence of experimental HM transmission in these studies, the detection of the pathogens in potential vectors nevertheless provides evidence of vector potential [43]. To underline the potential of Stomoxyx calcitrans as mechanical vectors, additional studies combining experimental studies and molecular methods such as quantification of porcine HMs in flies with different starvation intervals after blood meals should be considered in the future. The quantification of HM DNA in the flies examined revealed mean bacterial loads of 3.9 × 102 M. suis and 2.1 × 103 M. parvum for the total washing solutions and of 2.0 × 102 M. suis and 2.4 × 103 M. parvum in the crushed fly samples, respectively. To the best of our knowledge, this is the first molecular quantification study of porcine HMs in Stomoxys species. Similar quantities were found for M. wenyonii in Stomoxys samples from a Hungarian cattle herd, with a mean copy number of 1.0 × 102. The same study, however, revealed notably lower copy numbers for ‘Ca. M. haemobos’ (mean copy number of 1.0) in stable flies [40]. Based on their results, the authors suggested that the bacterial loads in hematophagous arthropods may be influenced by (1) the level of the bacteremia in livestock and, thus, the HM availability for potential blood-sucking vectors or (2) the differences in the transmission efficiency between HM species [40]. In the present study, HM blood levels of pigs at the time of fly sampling in the respective stables are unfortunately not available. Another factor influencing the bacterial load in the flies might be the decrease in bacteria with temporal distance from the blood meal. Prullage found that the transmission capability of Stomoxys calcitrans for M. suis is no longer present 24 h after the last blood meal [24]. Studies with other pathogens such as West Nile virus in stable flies also showed a rapid decrease in the number of flies containing the virus and the amount of virus within flies with distance to the blood meal [44]. Therefore, the bacterial loads detected in Stomoxys calcitrans, tested immediately after blood meals, may be different. In the present study, information on the time of the last blood meal is not available. However, several previous studies demonstrated that HMs can also be transmitted by hematophagous arthropods, even at low bacterial loads. In the case of M. ovis, it was shown that one infected red blood cell is sufficient for mechanical transmission. Consequently, M. ovis infection through natural vector-borne transmission can be initiated at a low level of parasitemia in sheep along with a high-level activity of blood-sucking insects [45].
In a second approach, we used high-throughput 16S rRNA amplicon sequencing as an alternative method for the investigation of Stomoxys calcitrans as a vector for HMs. This approach has the advantage over PCR-based methods in that not only known species can be detected but also those previously unknown. However, it was not possible to detect HM sequences by next generation amplicon sequencing in M. suis or M. parvum qPCR positive samples. Obviously, the HM loads in the Stomoxys samples investigated were too low. Rather than HM sequences, NGS results revealed mainly Morganella morganii ssp., Arthrobacter russicus and Phenylobacterium koreense sequences. The presence of various other bacteria in stable flies in combination with universal primers seems to result in superimposing the presence of the small amounts of HMs. This emphasizes that, for detecting small amounts of HMs, very specific methods, such as quantitative real-time PCR, are compulsory. The detection of the above-mentioned bacterial species was not surprising, since Schwarz et al. also detected Morganella morganii and Arthrobacter sp. by culture and matrix-assisted laser desorption ionization–time of flight mass spectrometry (MALDI-TOF MS) in fly samples out of the same Austrian pig stables (stables 1–20) [28].

5. Conclusions

The detection of hemoplasma DNA (M. parvum and M. suis) in stable flies indicates their potential to contribute to the spreading of HMs. Vector transmission through widespread Stomoxys calcitrans could possibly have an epidemiological impact, since it could allow for intra- and inter-herd HM spread. Since M. suis and M. parvum are highly prevalent in wild boars, HM vector transfer from the wildlife reservoir to pig farms should also be considered regarding biosafety measures. Moreover, Stomoxys calcitrans might be used to monitor the hemoplasma status in the stable, but the absence of HMs in stable flies does not allow for conclusions regarding the absence of HMs in pigs. Detected bacterial loads of HMs in stable flies are rather low. Therefore, precise and highly sensitive detection methods are required to prove the presence of HMs in stable flies. Further studies are called for in order to gain more knowledge on the role of Stomoxys calcitrans as a vector for HMs, such as experiments with larger numbers of flies and blood samples of HM positive and negative stables or quantification of HMs in flies with different starvation intervals after blood meals, to obtain information about persistence duration within the vector.

Author Contributions

Conceptualization, L.S., and J.S.; Investigation, M.A. and K.H.; Methodology, J.A., and J.S.; Writing—original draft, M.A., K.H., and J.A.; Writing—review and editing, M.R., L.E.H., J.S. and L.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Petri, F.A.M.; Sonalio, K.; de Souza Almeida, H.M.; Ferraz, M.E.S.; Storino, G.Y.; de Souza, M.R.; André, M.R.; de Oliveira, L.G. Porcine hemothropic mycoplasmas infection associated with productive impact in intensive pig production. Porcine Health Manag. 2020, 6, 33. [Google Scholar] [CrossRef]
  2. Bordin, L.C.; Gava, D.; Sonalio, K.; Mechler-Dreibi, M.L.; Zanella, J.R.C.; Morés, N.; de Oliveira, L.G.; Vaz, E.K. Investigation of hemotropic Mycoplasmas in fetuses and sows with reproductive failure. Vet. Anim. Sci. 2021, 12, 100175. [Google Scholar] [CrossRef]
  3. Messick, J.B. Hemotrophic mycoplasmas (hemoplasmas): A review and new insights into pathogenic potential. Vet. Clin. Pathol. 2004, 33, 2–13. [Google Scholar] [CrossRef]
  4. Santos, N.J.R.; Brito, D.R.B.; Abate, H.L.; Paixão, S.F.; Soares, E.D.S.; Vieira, T.S.W.J.; Garcia, J.L.; Vieira, R.F.C.; Vidotto, O. Hemotropic mycoplasmas infection in water buffaloes (Bubalus bubalis) from northeastern Brazil. Comp. Immunol. Microbiol. Infect. Dis. 2018, 56, 27–29. [Google Scholar] [CrossRef]
  5. Amer, M.M.; Soliman, A.M.; Do, T.; Hang, L.; El-Sayed, S.A.E.; Jaroszewski, J.; Mohanta, U.K.; Xuan, X. First molecular detection of hemotropic Mycoplasma spp. and molecular screening of other vector-borne pathogens in camels from the greater Cairo metropolitan area, Egypt. Trop. Biomed. 2024, 41, 412–421. [Google Scholar] [CrossRef]
  6. Willi, B.; Novacco, M.; Meli, M.; Wolf-Jäckel, G.; Boretti, F.; Wengi, N.; Lutz, H.; Hofmann-Lehmann, R. Haemotropic mycoplasmas of cats and dogs: Transmission, diagnosis, prevalence and importance in Europe. Schweiz. Arch. Tierheilkd. 2010, 152, 237–244. [Google Scholar] [CrossRef]
  7. Mascarelli, P.E.; Keel, M.K.; Yabsley, M.; Last, L.A.; Breitschwerdt, E.B.; Maggi, R.G. Hemotropic mycoplasmas in little brown bats (Myotis lucifugus). Parasit. Vectors 2014, 7, 117. [Google Scholar] [CrossRef]
  8. Westmoreland, L.S.H.; Stoskopf, M.K.; Maggi, R.G. Detection and prevalence of four different hemotropic Mycoplasma spp. in Eastern North Carolina American black bears (Ursus americanus). Comp. Immunol. Microbiol. Infect. Dis. 2017, 50, 106–109. [Google Scholar] [CrossRef]
  9. Maggi, R.G.; Chitwood, M.C.; Kennedy-Stoskopf, S.; DePerno, C.S. Novel hemotropic Mycoplasma species in white-tailed deer (Odocoileus virginianus). Comp. Immunol. Microbiol. Infect. Dis. 2013, 36, 607–611. [Google Scholar] [CrossRef]
  10. Stadler, J.; Jannasch, C.; Mack, S.L.; Dietz, S.; Zöls, S.; Ritzmann, M.; Hoelzle, K.; Hoelzle, L.E. Clinical and haematological characterisation of Mycoplasma suis infections in splenectomised and non-splenectomised pigs. Vet. Microbiol. 2014, 172, 294–300. [Google Scholar] [CrossRef]
  11. Henry, S.C. Clinical observations on eperythrozoonosis. J. Am. Vet. Med. Assoc. 1979, 174, 601–603. [Google Scholar] [CrossRef] [PubMed]
  12. do Nascimento, N.C.; dos Santos, A.P.; Chu, Y.; Guimaraes, A.M.S.; Baird, A.N.; Weil, A.B.; Messick, J.B. Microscopy and genomic analysis of Mycoplasma parvum strain Indiana. Vet. Res. 2014, 45, 86. [Google Scholar] [CrossRef] [PubMed]
  13. Thongmeesee, K.; Narapakdeesakul, D.; Kaewthamasorn, M.; Wattanaphansak, S.; Tiawsirisup, S. Characterization of hemotropic Mycoplasma (hemoplasma) isolates from Thai domestic pigs using the 23S ribosomal RNA gene. Acta Trop. 2025, 268, 107698. [Google Scholar] [CrossRef]
  14. Thongmeesee, K.; Kamkong, P.; Thanee, S.; Wattanapansak, S.; Kaewthamasorn, M.; Tiawsirisup, S. Molecular detection and genetic analysis of porcine haemoplasmas in commercial pig farms from Thailand reveal a putative novel species. Transbound. Emerg. Dis. 2022, 69, e2028–e2040. [Google Scholar] [CrossRef]
  15. Seo, M.-G.; Kwon, O.-D.; Kwak, D. Prevalence and phylogenetic analysis of hemoplasma species in domestic pigs in Korea. Parasit. Vectors 2019, 12, 378. [Google Scholar] [CrossRef]
  16. Fu, Y.; Shi, T.; Xu, L.; Wei, W.; Lu, F.; Zhang, X.; Yuan, X.; Li, J.; Lv, J.; Fang, W. Identification of a novel Hemoplasma species from pigs in Zhejiang province, China. J. Vet. Med. Sci. 2017, 79, 864–870. [Google Scholar] [CrossRef]
  17. Stadler, J.; Ade, J.; Ritzmann, M.; Hoelzle, K.; Hoelzle, L.E. Detection of a novel haemoplasma species in fattening pigs with skin alterations, fever and anaemia. Vet. Rec. 2020, 187, 66. [Google Scholar] [CrossRef]
  18. Thongmeesee, K.; Chonglomkrod, B.; Srisakdi, C.; Saributr, M.; Suksai, P.; Kamkong, P.; Tiawsirisup, S. Molecular detection of Mycoplasma wenyonii and its closely related hemotropic Mycoplasma sp. in blood-sucking flies from a buffalo farm in Chachoengsao province, Thailand. Acta Trop. 2022, 235, 106647. [Google Scholar] [CrossRef]
  19. Heinritzi, K. Untersuchungen zur Übertragbarkeit von Eperythrozoon suis. Tierarztl. Umsch. 1992, 47, 588–599. [Google Scholar]
  20. Hoelzle, L.E. Haemotrophic mycoplasmas: Recent advances in Mycoplasma suis. Vet. Microbiol. 2008, 130, 215–226. [Google Scholar] [CrossRef]
  21. Ade, J.; Ritzmann, M.; Wöstmann, C.; Eddicks, M.; Reese, S.; Hoelzle, K.; Hoelzle, L.E.; Stadler, J. Update on shedding and transmission routes of porcine haemotrophic mycoplasmas in naturally and experimentally infected pigs. Porcine Health Manag. 2021, 7, 49. [Google Scholar] [CrossRef] [PubMed]
  22. Henderson, J.P.; O’Hagan, J.; Hawe, S.M.; Pratt, M.C. Anaemia and low viability in piglets infected with Eperythrozoon suis. Vet. Rec. 1997, 140, 144–146. [Google Scholar] [CrossRef] [PubMed]
  23. Acosta, D.B.; Ruiz, M.; Sanchez, J.P. First molecular detection of Mycoplasma suis in the pig louse Haematopinus suis (Phthiraptera: Anoplura) from Argentina. Acta Trop. 2019, 194, 165–168. [Google Scholar] [CrossRef] [PubMed]
  24. Prullage, J.B.; Williams, R.E.; Gaafar, S.M. On the transmissibility of Eperythrozoon suis by Stomoxys calcitrans and Aedes aegypti. Vet. Parasitol. 1993, 50, 125–135. [Google Scholar] [CrossRef]
  25. Sprygin, A.; Pestova, Y.; Wallace, D.B.; Tuppurainen, E.; Kononov, A.V. Transmission of lumpy skin disease virus: A short review. Virus Res. 2019, 269, 197637. [Google Scholar] [CrossRef]
  26. Mellor, P.S.; Kitching, R.P.; Wilkinson, P.J. Mechanical transmission of capripox virus and African swine fever virus by Stomoxys calcitrans. Res. Vet. Sci. 1987, 43, 109–112. [Google Scholar] [CrossRef]
  27. Schuberg, A.; Böing, W. Über die Übertragung von Krankheiten durch einheimische stechende Insekten (On the transmission of diseases by indigenous biting insects). Arb. Kaiserl. Gesundh. 1914, 48, 491–512. [Google Scholar]
  28. Schwarz, L.; Strauss, A.; Loncaric, I.; Spergser, J.; Auer, A.; Rümenapf, T.; Ladinig, A. The Stable Fly (Stomoxys calcitrans) as a Possible Vector Transmitting Pathogens in Austrian Pig Farms. Microorganisms 2020, 8, 1476. [Google Scholar] [CrossRef]
  29. Moon, R.D. Muscid flies (Muscidae). In Medical and Veterinary Entomology, 3rd ed.; Mullen, G.R., Durden, L.A., Eds.; Academic Press an Imprint of Elsevier: London, UK, 2019; ISBN 0128140445. [Google Scholar]
  30. Paslaru, A.I.; Verhulst, N.O.; Maurer, L.M.; Brendle, A.; Pauli, N.; Vögtlin, A.; Renzullo, S.; Ruedin, Y.; Hoffmann, B.; Torgerson, P.R.; et al. Potential mechanical transmission of Lumpy skin disease virus (LSDV) by the stable fly (Stomoxys calcitrans) through regurgitation and defecation. Curr. Res. Insect Sci. 2021, 1, 100007. [Google Scholar] [CrossRef]
  31. Baldacchino, F.; Muenworn, V.; Desquesnes, M.; Desoli, F.; Charoenviriyaphap, T.; Duvallet, G. Transmission of pathogens by Stomoxys flies (Diptera, Muscidae): A review. Parasite 2013, 20, 26. [Google Scholar] [CrossRef]
  32. Schofield, S.; Torr, S.J. A comparison of the feeding behaviour of tsetse and stable flies. Med. Vet. Entomol. 2002, 16, 177–185. [Google Scholar] [CrossRef] [PubMed]
  33. Ade, J.; Hoelzle, K.; Stadler, J.; Ritzmann, M.; Hoelzle, L.E. Occurrence of Mycoplasma parvum in German Pigs of Different Age Groups Using a Novel Quantitative Real-Time PCR Assay. Pathogens 2022, 11, 1374. [Google Scholar] [CrossRef] [PubMed]
  34. Hoelzle, L.E.; Helbling, M.; Hoelzle, K.; Ritzmann, M.; Heinritzi, K.; Wittenbrink, M.M. First LightCycler real-time PCR assay for the quantitative detection of Mycoplasma suis in clinical samples. J. Microbiol. Methods 2007, 70, 346–354. [Google Scholar] [CrossRef]
  35. Ade, J.; Stadler, J.; Ritzmann, M.; Zübert, C.; Hoelzle, K.; Hoelzle, L.E. Occurrence of ‘Candidatus Mycoplasma haemosuis’ in fattening pigs, sows and piglets in Germany using a novel gap-based quantitative real-time PCR assay. BMC Vet. Res. 2022, 18, 40. [Google Scholar] [CrossRef]
  36. Microsynth, A.G. User Guide Amplicon Metagenomics. Available online: https://www.microsynth.com/files/Inhalte/PDFs/Next%20Generation%20Sequencing/UserGuide_AmpliconMetagenomics.pdf (accessed on 6 May 2025).
  37. Zumpt, F. The Stomoxyine Biting Flies of the World. Diptera: Muscidae. Taxonomy, Biology, Economic Importance and Control Measures; G. Fischer: Stuttgart, Germany, 1973. [Google Scholar]
  38. Butler, J.F.; Kloft, W.J.; DuBose, L.A.; Kloft, E.S. Recontamination of food after feeding a 32P food source to biting muscidae. J. Med. Entomol. 1977, 13, 567–571. [Google Scholar] [CrossRef]
  39. Song, Q.; Zhang, W.; Song, W.; Liu, Z.; Khan, M.K.; He, L.; Fang, R.; Li, P.; Zhou, Y.; Hu, M.; et al. Seroprevalence and risk factors of Mycoplasma suis infection in pig farms in central China. Prev. Vet. Med. 2014, 117, 215–221. [Google Scholar] [CrossRef]
  40. Hornok, S.; Micsutka, A.; Meli, M.L.; Lutz, H.; Hofmann-Lehmann, R. Molecular investigation of transplacental and vector-borne transmission of bovine haemoplasmas. Vet. Microbiol. 2011, 152, 411–414. [Google Scholar] [CrossRef]
  41. Overas, J. Studies on Eperythrozoon ovis infection in sheep. Acta Vet. Scand. 1969, 28, 148. [Google Scholar]
  42. Nonaka, N.; Thacker, B.J.; van Schillhorn Veen, T.W.; Bull, R.W. In vitro maintenance of Eperythrozoon suis. Vet. Parasitol. 1996, 61, 181–199. [Google Scholar] [CrossRef]
  43. Arendt, M.; Stadler, J.; Ritzmann, M.; Ade, J.; Hoelzle, K.; Hoelzle, L.E. Hemotrophic Mycoplasmas-Vector Transmission in Livestock. Microorganisms 2024, 12, 1278. [Google Scholar] [CrossRef]
  44. Doyle, M.S.; Swope, B.N.; Hogsette, J.A.; Burkhalter, K.L.; Savage, H.M.; Nasci, R.S. Vector competence of the stable fly (Diptera: Muscidae) for West Nile virus. J. Med. Entomol. 2011, 48, 656–668. [Google Scholar] [CrossRef] [PubMed]
  45. Mason, R.W.; Statham, P. The determination of the level of Eperythrozoon ovis parasitaemia in chronically infected sheep and its significance to the spread of infection. Aust. Vet. J. 1991, 68, 115–116. [Google Scholar] [CrossRef]
Table 1. Results of M. suis-specific real-time PCRs.
Table 1. Results of M. suis-specific real-time PCRs.
Sample SourceWash Solution Fly
Positive/All TestedCopy Number in Total Positive Wash Solution Positive/All TestedCopy Number in Positive Fly
stable 40/3n.d.1/32.9 × 101
stable 61/38.4 × 1021/33.8 × 102
stable 71/37.7 × 1021/38.0 × 101
stable 80/3n.d.1/32.6 × 102
stable 131/31.1 × 1031/32.5 × 102
stable 161/31.1 × 1030/3n.d.
Table 2. Results of M. parvum-specific real-time PCRs.
Table 2. Results of M. parvum-specific real-time PCRs.
Sample SourceWash Solution Fly
Positive/All TestedCopy Number in Total Positive Wash SolutionPositive/All TestedCopy Number in Positive Fly
stable 61/31.8 × 1030/3n.d.
stable 71/31.7 × 1030/3n.d.
stable 82/37.6 × 1020/3n.d.
1.8 × 103
stable 91/39.0 × 1020/3n.d.
stable 101/31.6 × 1030/3n.d.
stable 140/3n.d.1/33.3 × 103
stable 151/33.4 × 1020/3n.d.
stable 160/3n.d.1/31.9 × 103
stable 170/3n.d.1/31.9 × 103
stable 181/34.6 × 1030/3n.d.
stable 191/35.9 × 1031/33.9 × 103
stable 220/6n.d.1/61.2 × 103
Table 3. Results of next generation sequencing.
Table 3. Results of next generation sequencing.
Operational Taxonomic Unit (OUT)BacteriaPresence (%)
ID Sample 1Sample 2
1Morganella morganii subsp. morganii30.10.5043
2Morganella morganii subsp. morganii13.940.1441
3Morganella morganii subsp. sibonii13.930.2161
4Morganella morganii subsp. morganii13.820.07205
5Morganella morganii subsp. sibonii13.330.07205
6Morganella morganii subsp. sibonii13.260.2161
7Arthrobacter russicus1.58296.61
8Phenylobacterium koreense0.025612.161
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Arendt, M.; Hoelzle, K.; Stadler, J.; Ritzmann, M.; Ade, J.; Hoelzle, L.E.; Schwarz, L. Occurrence and Quantification of Porcine Hemotrophic Mycoplasmas in Blood-Sucking Stomoxys calcitrans. Microorganisms 2025, 13, 1607. https://doi.org/10.3390/microorganisms13071607

AMA Style

Arendt M, Hoelzle K, Stadler J, Ritzmann M, Ade J, Hoelzle LE, Schwarz L. Occurrence and Quantification of Porcine Hemotrophic Mycoplasmas in Blood-Sucking Stomoxys calcitrans. Microorganisms. 2025; 13(7):1607. https://doi.org/10.3390/microorganisms13071607

Chicago/Turabian Style

Arendt, Mareike, Katharina Hoelzle, Julia Stadler, Mathias Ritzmann, Julia Ade, Ludwig E. Hoelzle, and Lukas Schwarz. 2025. "Occurrence and Quantification of Porcine Hemotrophic Mycoplasmas in Blood-Sucking Stomoxys calcitrans" Microorganisms 13, no. 7: 1607. https://doi.org/10.3390/microorganisms13071607

APA Style

Arendt, M., Hoelzle, K., Stadler, J., Ritzmann, M., Ade, J., Hoelzle, L. E., & Schwarz, L. (2025). Occurrence and Quantification of Porcine Hemotrophic Mycoplasmas in Blood-Sucking Stomoxys calcitrans. Microorganisms, 13(7), 1607. https://doi.org/10.3390/microorganisms13071607

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop