Checklist of Medico-Veterinary Important Biting Flies (Ceratopogonidae, Hippoboscidae, Phlebotominae, Simuliidae, Stomoxyini, and Tabanidae) and Their Associated Pathogens and Hosts in Maghreb

Abstract

Biting flies are hematophagous dipterans belonging to various taxonomic groups, such as the Hippoboscidae, Ceratopogonidae, Simuliidae, Tabanidae, Muscidae, and Psychodidae families, some of which have significant medical and veterinary importance. They can host and spread various infections to humans and livestock and cause allergic reactions with their saliva. Several species of different families are present in the western Mediterranean region, with new species gradually being discovered. This study focuses on the brachyceran and the nematoceran species; it provides a systematic review listing all reported taxa of biting flies in the Maghreb countries (Algeria, Morocco, and Tunisia). Additionally, the study includes a geo-historical reconstruction of distribution maps for species of epidemiological importance. The associated pathogens and hosts are also included in the checklists, alongside information on the biology and ecology of these parasitic arthropods, to offer a comprehensive overview of the state of dipteran-borne disease surveillance in North African countries. Overall, this work could serve as an exhaustive reference for entomologists and breeders participating in controlling biting fly and midge populations, whether from a technical or research perspective.

1. Background

The Diptera order is among the world’s most significant, varied, and well-known insect orders. This order name significates “two-winged”, which describes a substantially altered and reduced rear pair of wings [1]. Nearly 160,000 recognized taxa are known, divided into approximately 10,000 genera and 150 families [2]. The number of species continues to increase, as new species are regularly identified [3].

Dipteran species are classified based on morphological criteria, such as wing morphology, wing vein number, abdominal patterns, chaetotaxy, genitalia, and mouthpart structure [4]. However, advanced tools such as ultramicroscopy and molecular technologies in entomology, such as PCR and sequencing, RFLP PCR, DNA barcoding, and MALDI-TOF MS profiling, have become essential for dipteran species identification [5,6,7].

Recent molecular tools enable researchers to precisely identify and deeply understand their genetic bases, among them, their feeding behavior. It is known that biting flies adopt hematophagy as a principal feeding behavior; this hematophagy can be common to both sexes in certain dipteran taxa such as Stomoxyini, and it can result from dimorphic features separating males and females in many other taxa, including Culicidae, Ceratopgonidae, Simulidae, Tabanidae, and phlebotomine. In these cases, only females are hematophagous, every so often requiring a blood meal or more to lay their eggs. Males are usually more adapted to feed on nectar, honeydew, and other plant fluids [8].

The blood-sucking behavior of dipterans on its own is a serious medical and economic issue. When a dipteran pierces the skin, hosts often experience pain, and in some cases, an excessive allergic reaction may occur, characterized by erythema, itching, and swelling. Though rare, anaphylactic shock can also develop in extreme cases [9]. Furthermore, other dipteran bites, such as tabanids, represent an important source of disturbance, particularly for harvesting crews and farm laborers in areas of intensive agriculture [10]. Other dipterans, such as stable flies, have detrimental effects on livestock health, impacting thriftiness, animal weight gain, and milk production. This is particularly significant on farms, where multiple species with alternating daily activity patterns coexist. Some species may be active throughout the day, while others are primarily active in the morning and evening [10,11,12]. Others, such as biting midges (Ceratopogonidae), have been identified as significant sources of nuisance at major tourist attractions, limiting recreational activities, particularly in areas with high population densities of these species [13,14,15].

Several diseases have been associated with different species of parasitic dipterans, particularly in the Maghreb region. These include leishmaniasis, phlebovirus infections, trypanosomiasis, leucocytozoonosis, bluetongue (catarrhal fever), bartonellosis, rickettsiosis, and many others (Supplementary Materials).

The North African region, spanning from Morocco to Tunisia, has a unique demographic profile shaped by its location as a transitional zone between the Palearctic and Afrotropical climatic regions. Its high biodiversity, varied landscapes, and coastal areas contribute to the region’s vulnerability to dipteran-borne diseases like trypanosomiasis, arbovirus, leishmaniasis, and other diseases [16].

Taking the example of leishmaniases, a widespread disease in the Mediterranean, especially the Maghreb region, with 44,050 cases, Algeria leads the globe in cases of Cutaneous leishmaniasis, second after Afghanistan, Tunisia (7631), and Morocco (3430). Although they are less common, Morocco leads the world in visceral leishmaniases with 152 cases, followed by Algeria with 111 and Tunisia with 89 between 2004 and 2008 [17].

A dense population, rapid urbanization, and extensive agricultural activities increase human exposure to dipteran vectors, while climate change exacerbates the spread of vector-borne diseases. The Maghreb’s ecological diversity, ranging from arid deserts to fertile plains, creates varied habitats for human populations and disease-carrying species, enhancing susceptibility to epidemics [18,19].

Distribution maps of the Euro-Mediterranean biting fly species, mainly those of the Aedes, Anopheles, and Culex genera, which include Algeria, Egypt, Libya, Morocco, and Tunisia, have been released. Afterward, mosquito population inventories were assessed in various geographical areas of Algeria [20,21]. More recently, the status of mosquito-borne diseases and their vectors in North Africa was updated and reviewed [22]. However, the status of Brachyceran and Nematoceran biting flies in Algeria and North Africa remains limited and poorly investigated.

Therefore, understanding the diversity and ecology of biting flies and midges is paramount for effective disease management and control strategies. Given the complexity of these insects’ life cycles, ecological preferences, and vector competencies, a comprehensive review of the main biting species is mandatory. This review aims to provide an update on biting-fly-borne pathogens and their vectors in the Maghreb region (Algeria, Tunisia, and Morocco) by collecting scientific data about dipterans of medical and veterinary interest and their hosts. It also aims to build distributional maps of the main biting fly species across the Maghreb. Consequently, a good understanding of the epidemiology of the diseases transmitted by these flies is required to implement appropriate control strategies.

2. Methods

A systematic literature review was conducted in French and English using different search engines, including Google Scholar, Internet Archive, Pubmed, Semantic Scholars, and Scopus. Different combinations of English keywords were used for each taxon and then repeated in French. Some of these keywords included the following:

‘Ceratopongidae or Biting midges or Culicoides’ and ‘Algeria or Morocco or Tunisia’, or ‘Ceratopongidae or Biting midges or Culicoides’ and ‘Pathogens or viruses or Bluetongue or Onchocerca or Leishmania’ and ‘Algeria or Morocco or Tunisia’.

‘Phlebotomes or sand flies’ and ‘Algeria or Morocco or Tunisia’, or ‘Phlebotomes or sand flies’ and ‘Leishmania or pathogens or phleboviruses’ and ‘Algeria or Morocco or Tunisia’. ‘Simuliids or black flies’ and ‘Algeria or Morocco or Tunisia’, or ‘simuliids or black flies’ and ‘Leucocytozoon or Onchocerca or pathogens or phleboviruses’ and ‘Algeria or Morocco or Tunisia’.

The collected data included published (peer-reviewed papers) and grey literature (unpublished PhD theses, book chapters, and Pasteur Institute archive notes). Additionally, three authors were contacted via email to request raw data and confirm their identification, but no reply was received.

Geographical coordinates were extracted from the consulted references to establish distributional maps of the dipterans of interest using QGIS software (Geographic Information System, Version 3.30). When the reference lacked coordinates but mentioned the governorate or gave a complete location description, the GPS coordinates were estimated based on the description of the core text.

Some references from the French colonial era, including archives and annals from the Pasteur Institute in the three countries, contained the ancient designations of locations, which required historical cartographic research to establish their current names and, thus, their coordinates.

Given the presence of various synonyms in the nomenclature of different dipteran taxa in the Maghreb (e.g., Ceratopogonidae, Phlebotominae, Simuliidae, and Tabanidae), cross-referencing the literature was essential. This step was carried out by consulting relevant catalogs to ensure accurate taxonomic confirmation, such as the Catalog of the Biting Midges of the World (Diptera: Ceratopogonidae; [23]), Catalogue of the Diptera (Insecta) of Morocco [24], world’s simuliids comprehensive revision of the taxonomic and geographical inventory [25], and the Synoptic Catalogue of world’s Tabanids [26].

3. Main Biting Fly Groups of Medical and Veterinary Importance in the Maghreb

3.1. Stomoxyini

The Stomoxys genus belongs to the Stomoxyini tribe, from the Muscinae subfamily and Muscidae family, which appeared about 30 million years ago in the eastern region and includes 18 species, 17 of which have a tropical distribution [27]. They are obligate blood-suckers, and several species are often considered to be important economic pests for livestock and other warm-blooded animals [27].

Commonly known as “stable flies”, Stomoxys calcitrans (Linnaeus, 1758) is the only cosmopolitan species and the most well-studied member of the Stomoxyini tribe worldwide [28]. Haematobia irritans (Linnaeus, 1758), found in the Maghreb, is another species of medical and veterinary importance within this group. However, to the best of our knowledge, studies focused on stomoxyini flies in North Africa remain scarce.

Stable flies can be found breeding in dead vegetation near flooded areas and beaches, in grass clippings, at the base of silage heaps, in improperly managed compost piles, and in hay bales [29]. Eggs are laid in moist organic materials such as litter, manure, and straw and hatch at an ideal temperature of 32 °C and a 90% humidity [30].

The life cycle of stable flies consists of the following four stages: egg, larva, pupa, and adult. Female stable flies lay their eggs in moist, decaying organic matter, often contaminated with animal waste [29,31]. The eggs hatch within 12–24 h into small larvae that develop in moist, fermenting plant materials, such as silage or rotting vegetables, and depend on the microbial communities within these substrates for growth [32]. The larvae feed on the fermenting organic matter, progressing through three larval instars over approximately 12–13 days, depending on environmental conditions such as temperature, substrate quality, and larval density [33].

As they progress through these stages, the larvae molt between each instar, eventually reaching the third instar, forming a puparium [34,35,36]. This pupal stage lasts approximately seven days, after which the adult fly emerges. The complete development from egg to adult generally takes around 20 days under optimal conditions (e.g., 27 °C). However, the development time can vary with the breeding site’s temperature, substrate, and larval density changes [34,36].

Once emerged, adults of both sexes are aggressive and persistent blood-feeders. They can even attack and feed on humans when their preferred hosts are absent [37]. However, it has additionally been proven that S. calcitrans and S. niger (Macquart, 1851) are nectarivorous or ripe fruit feeders [36].

These feeding habits of the Stomoxyini govern their interactions with their hosts, as well as with pathogens. Therefore, stable flies affect livestock productivity and human livelihood [38]. Because of their bites, these insects are very annoying to livestock. They can induce skin lesions and defensive movements of the head, ears, and legs that stress animals, resulting in energy loss and a substantial decrease in milk and meat production. For instance, in the US, economic losses due to stable fly bites are estimated at USD 1 billion [37].

They also play a significant role as mechanical vectors of various pathogens, including bacteria, viruses, and protozoa, especially S. calcitrans, passively transmitting diseases to livestock and humans. Although the stable fly is primarily known for its painful bite and nuisance behavior, its involvement in pathogen transmission is being increasingly recognized. These pathogens have been detected or experimentally confirmed in Stomoxys spp., contributing to their relevance in veterinary and medical entomology. For instance, S. calcitrans has been suggested as a potential mechanical vector of the protozoan Besnoitia besnoiti [39]. Other pathogens, such as viruses, are transmitted either by contaminated mouthparts, the regurgitation of the blood meal, or defecation [40].

Trypanosoma evansi, the surra agent, has been associated with various Stomoxys species. In Tunisia, T. evansi was detected in Stomoxys sp., with Canis familiaris as the affected host [41]. Additionally, in controlled experiments, S. niger niger and S. taeniatus transmitted T. evansi to laboratory mice, highlighting their potential in the spread of surra in susceptible hosts [42,43]. Other trypanosome species, such as T. brucei and T. congolense, were found to be transmitted by S. niger bileneatus, S. varipes, and S. pallidus, especially to BALB/c mice in experimental settings [42,43,44]. In addition to T. brucei, the mechanical transmission of other Nagana-responsible agents such as T. vivax by S. n. bileneatus, S. n. niger, and Haematobosca squalida has been previously recorded in Africa alongside the mechanical transmission of T. simiae affecting swine species. This protozoan was found to be transported by S. omega and S. n. niger [42,44].

Beyond protozoans, other disease agents, such as viruses, can be spread mechanically by stomoxes. For instance, S. calcitrans has been experimentally linked to the equine infectious anemia virus (EIAV) spread to Equus caballus [45]. On the other hand, there is experimental evidence of African swine fever virus (ASFV) transmission by S. calcitrans to swine. In contrast, West Nile fever virus (WNFV) was detected in S. calcitrans and was associated with the American white pelican Pelecanus erythrorhynchos [46,47]. Furthermore, the experimental infection of golden hamsters (Mesocricetus auratus) with Rift Valley fever virus (RVFV) through S. calcitrans indicates the potential for zoonotic disease spread [48]. S. calcitrans has also been suspected in several studies to be the potential mechanical vector of LSD, Lympy Skin Disease [49,50,51].

Additionally, S. calcitrans is implicated in the mechanical transmission of bacterial pathogens, including food-borne disease agents such as Enterobacter sakazakii, which can cause severe infections in newborns [52]. Other bacterial agents associated with S. calcitrans include Bacillus anthracis (anthrax) and Anaplasma marginale, although definitive host data are lacking. Furthermore, Coxiella brunetii, the causative agent of Q fever, has also been found in S. calcitrans, with transmission observed in cattle and elk in North America [50,53,54].

Traversa, Otranto [55] demonstrated that S. calcitrans can serve as an intermediate host, allowing for the development of Habronema microstoma’s lifecycle, making this one of the few pathogens that stable flies can biologically transmit. Considering their role (which is mainly mechanical) in the spread of these pathogens emphasizes the need for profound studies targeting these groups of dipterans, especially with the epizootic nature of LSD that Algeria is facing during the current year.

3.2. Hippoboscidae

Flies belonging to the Hippoboscidae family are commonly known as “louse flies”, “forest flies”, or “keds” [56]. These flies are considered to be parasites of wild and domestic animals and occasionally humans [56,57]. The Hippoboscidae family members exhibit potent characteristics, with some species possessing reasonably robust, permanent wings, while others lose their wings upon reaching their hosts with their flattened bodies and powerful mouthparts. Both males and females are solenophages [58,59].

In addition to their morphological traits, members of the Hippoboscidae family exhibit a specific reproductive behavior called adenotrophic viviparity, a mode of reproduction wherein larvae grow inside the female and are fed by secretory glands. The uterus and the female accessory gland expand and adapt to offer the synthesis and transport of nutrients and a habitat for developing larvae [56,60]. After mating, a single egg is passed to the uterus, where the embryo feeds on the yolk until hatching. The first and second larval instars remain in the uterus, where a pair of intra-uterine secretory milk glands nourish them. This mechanism is similar to the one observed in tsetse flies. During this feeding process, endosymbionts are passed from the female to the offspring [56].

Upon reaching the third instar, the larva stops to feed, and parturition occurs, leading to the larva’s emergence, during which, the integument hardens under hormone effects, forming the puparium, a rigid shell inside of which flies and other insects pupate [61]. Opposingly, some species’ internal pupal transformations are initiated in the uterus. Puparia are usually deposited near the host, mainly in the bedding area, which is a suitable location [56,61]. However, the sheep ked is exceptional, because females glue the puparium to the host’s fleece. After that, pupal development occurs, and an adult individual emerges within several weeks or months, depending on the species and temperature [56].

Understanding the complex lifecycle of the Hippoboscidae family provides a foundation for exploring their diverse classification. Eleven genera of keds are encountered in the Maghreb region, out of which, 20 species can be enumerated (

Table 1. Checklist of keds morphologically identified in the Maghreb region (Morocco, Algeria, and Tunisia).

Genus	Taxon	Taxonomy	Country
Crataerina	Crataerina. acutipennis	Austen, 1926	M
	Crataerina pallida	Latreille, 1811	M
Hippobosca	Hippobosca camelina	Leach, 1817	M, A, T
	Hippobosca equina	Linnaeus, 1758	M, A, T
	Hippobosca fulva	Austen, 1912	M
	Hippobosca longipennis	Fabricius, 1805	M, A, T
	Hippobosca variegata	Megerle, 1803	M
Icosta	Icosta minor	Bigot, 1858	M
Ornithoica	Ornithoica turdi	Olivier in Latreille, 1811	M
Ornithomyia	Ornithomyia avicularia	Linnaeus, 1758	M
	Ornithomyia fringilina	Curtis, 1836	A
	Ornithomyia biloba	Dufor, 1827	M, A
Ornithophila	Ornithophila gestroi	Rondani, 1878	M, A, T
	Ornithophila metallica	Schiner, 1864	M, A
Pseudolynchia	Pseudolynchia canariensis	Macquart, 1839	M, A
Stenepteryx	Stenepteryx hirundinis	Linnaeus, 1758	M
Lipoptena	Lipoptena capreoli	Rondani, 1878	M
	Lipoptena cervi	Linnaeus, 1758	A
Melophagus	Melophagus ovinus	Linnaeus, 1758	M, A, T
Lynchia	Lynchia pilosa	Macquart, 1843	M

A Algeria, M Morocco, T Tunisia.

). Species within the Hippobosca genus are known for their low host specificity [57], making them both biological and mechanical vectors of various zoonotic diseases affecting different animal species, including the dromedary camel (Camelus dromedarius), which has been found to be infested by T. evansi and T. vivax, both mechanically vectorized by H. camelina, as well as other Trypanosoma and Anaplasma vectorized by H. equina [62].

Another interesting genus of keds is Melophagus, which includes four parasitic species known as “louse flies” or “sheep keds”, which are as follows: M. antilopes, M. rupicaprinus, M. kamtshaticus, and M. ovinus. M. kamtshaticus is an obligate parasite of the bighorn sheep Ovis canadensis endemic to the Kamtchatka peninsula [63,64].

The domestic horse (Equus caballus) has been found to be infected by Bartonella melophagi, Bartonella chomelii, and various species of Anaplasma, all detected in its related keds, H. equina [65]. Cattle (Bos taurus) have also been found to be infected by different Trypanosoma and Bartonella species, including B. chomelii transmitted by H. equina and H. longipennis [66]. The spectrum of microorganisms transmitted by keds extends to wild animals such as the red deer, in which the North African-endemic subspecies (Cervus elaphus barbarus) is found to harbor various microorganisms, including Bartonella, Trypanosoma species, Rickettsia Helvetica, and filarioids. All were detected in its related keds, Lipoptena cervi. These findings suggest that wild hosts and their specific keds may maintain the microorganisms in the environment through the persistence of their sylvatic cycle [67,68].

In addition to the information provided earlier, the wingless fly genus under consideration includes one species of medical and veterinary interest, M. ovinus Linnaeus 1758. [63,69]. This species primarily parasitizes domestic sheep (Ovis aries), the main host throughout its life cycle. Nevertheless, reports have indicated its presence on other domestic and wild animals, including goats (Capra hircus), rabbits (Oryctolagus cuniculus), dogs (Canis familiaris), European bison (Bison bonasus), red foxes (Vulpes vulpes), and Humans [70,71].

Examples of possible pathogen transmission by M. ovinus (Figure 1a) include the suggested transmission of the border disease virus in small ruminants [72] and the mechanical transmission of the bluetongue virus [73]. Moreover, several studies from Europe and North Africa have reported the detection of different pathogens in sheep keds, including Anaplasma ovis, Anaplasma phagocytophilum, Bartonella. melophagi, and Borrelia burgdorferi. Additionally, other unidentified species from genera such as Bartonella, Borrelia, Rickettsia, Anaplasma, Acinetobacter, and Trypanosoma have also been documented [65,67,68].

The range of ked species of veterinary importance (Figure 2a) spans Morocco’s Atlas and peri-Atlas regions, with H. equina showing a particular affinity for Atlantic coastal areas. In contrast, the range of H. camelina extends across the western high plateaus and coastal regions of Algeria and the Dahar Mountains in southern Tunisia. Moreover, H. equina exhibits a unique pattern of range, stretching from near the western coast to the extreme eastern coast of Algeria. This species has also been recovered from the north of the Tunisian coastal governorate Mednine and Djerba Island, suggesting that its distribution pattern may be associated with low-altitude, humid, and warm Mediterranean regions.

Conversely, M. ovinus is typically found in Morocco’s Middle Atlas and Atlas regions, the Blidean Atlas and Djurdjura Mountains in Algeria, and the Dahar Mountains in southern Tunisia. This distribution pattern, previously reported by Kumsa, Parola [74], suggests that M. ovinus is limited to higher altitudes in tropical regions, driven more by altitude and environmental factors than host-dependent factors.

3.3. Ceratopogonidae

Biting midges (Ceratopogonidae) are cosmopolitan bloodsucking flies. Some of their names are punkies, derived from the Indian American word “Punkwa” and “Brulôt”, derived from the French verb “burler” (to burn), the first meaning ashlike, referring to their appearance while biting. In contrast, the second is more linked to the unpleasant burning sensation related to their bites [23,75].

This family includes approximately 6206 extant species listed in four subfamilies that are, to date, as follows: Leptoconopinae, Forcipomyiinae, Dasyheleinae, and Ceratopogoninae. All of them are hematophagous, except the Dasyheleinae group members [23].

Regarding their lifecycle, one week after their emergence, autogenous females oviposit a batch of eggs, while anautogenous females lay their eggs directly after their first blood meal. A second gonotrophic cycle is possible for females who can obtain a second blood meal, but seldom a third one or more outside laboratory experimentations [76,77]. Most species are multivoltine, producing at least two generations yearly [75]. It is noteworthy that their batches of eggs are usually deposited on moist substrates that constitute the breeding sites for Ceratopogonids. The larvae develop in these moist substrates and are found in aquatic and semiaquatic environments across diverse biomes, from tropical regions to tundra. Some species, such as Leptoconops larvae, are found in specific environments like clay-like soils in arid regions, tidal margins, and beaches [75], while larvae of the Forcipomyia genus are often found in shallow water and rotting wood. As for Culicoides larvae, they can be encountered in a variety of habitats, including freshwater swamps, the shallow margins of ponds, streams, and rivers, tidal margins, rotting wood and cacti, and animal manure surrounding leaking water [75,78].

Midges’ larvae are omnivorous, and many species are predators of rotifers, oligochaetes, protozoans, nematodes, and the immature stages of other invertebrates, including other ceratopogonid larvae [79,80,81]. Their feeding habits also include bacteria, fungi, microalgae, and diatoms [75,80]. The long and slender larvae lack spiracles, which makes their respiration cuticular. For some species, this is facilitated by the hygroscopic fluid of their secretory setae [82,83].

Depending on the species, their development occurs through four instars from over two weeks up to more than one year. During hot summers, they tend to become dormant [75].

The larval stage ends with pupation near the substrate’s surface. For tree hole species, the pupae typically float on the water’s surface, loosely adhering to the sides of the tree cavity. In most species, the buoyancy of the pupae during this life stage is maintained by an air pocket beneath the wings [23,75].

Males typically emerge before females at the end of the cycle. As their sperm matures within 24 h, they are ready to mate once the females emerge. Some aspects of their reproductive behavior resemble those observed in simuliids, including the swarming of males near water surfaces or breeding sites. Females of the same species passing through the swarm are recognized by their wing beat frequency and released pheromones [23,79]. Although both males and females feed on nectars, only females are hematophagous, because the blood acquired during feeding will support oogenesis [23].

During the last century, several species have been described from the Maghreb region (Algeria, Morocco, and Tunisia), and many of them were new to this region. The following eleven genera belonging to four subfamilies exist in this region: Alluaudomyia, Atrichopogon, Bezzia, Brachypogon, Culicoides, Dasyhelea, Forcipomyia, Leptoconops, Monohelea, Palpomyia, and Parabezzia (Supplementary Materials). However, to our knowledge, only the Culicoides and Forcipomyia genera are of veterinary and medical importance in the region, as they are vectors of several pathogens. The 114 species (83 Culicoides spp. and 33 Forcipomyia spp.) found in the Maghreb are listed in

Table 2. Checklist of midge species of Culicoides and Forcipomyia genera in the Maghreb with their respective identification methods.

Taxon	Taxonomy	Country	Identification
Culicoides achrayi	Kettle & Lawson, 1955	A	Morphological
Culicoides albicans	Winnertz, 1852	A	Morphological
Culicoides algeriensis	Clastrier, 1957	A	Morphological
Culicoides azerbajdzhanicus	Dzhafarov, 1962	A, M	Morphological
Culicoides badooshensis	Khalaf, 1961	M	Morphological
Culicoides begueti	Clastrier, 1957	A	Morphological
Culicoides calloti	Kremer, Delécolle, Bailly-Choumara & Chaker, 1979	M	Morphological
Culicoides cataneii	Clastrier, 1957	A, M, T	Morphological + PCR and sequencing
Culicoides chiopterus	Meigen, 1830	A	Morphological
Culicoides circumscriptus	Kiefer, 1918	A, M, T	Morphological + PCR and sequencing
Culicoides clastrieri	Callot, Kremer & Deduit, 1962	A, M	Morphological
Culicoides corsicus	Kremer, Leberre & Beaucournu-Saguez, 1971	A, T	Morphological
Culicoides derisor	Callot & Kremer, 1965	M	Morphological
Culicoides dewulf	Goetghebuer, 1936	A	Morphological
Culicoides duddingstoni	Kettle & Lawson, 1955	A	Morphological
Culicoides duddingstoni	Kettle & Lawson, 1955	M	Morphological
Culicoides dzhafarovi	Remm, 1967	A, M	Morphological
Culicoides faghihi	Navai, 1971	A, M	Morphological
Culicoides fagineus	Edwards, 1939	A, M	Morphological
Culicoides fascipennis	Staeger, 1839	A, T	Morphological
Culicoides festivipennis	Kiefer, 1914	A, M	Morphological
Culicoides foleyi	Kiefer, 1922	A	Morphological
Culicoides gejgelensis	Dzhafarov, 1964	A, M, T	Morphological + PCR and sequencing
Culicoides griseidorsum	Kiefer, 1918	A, T	Morphological
Culicoides grisescens	Edwards, 1939	A	Morphological
Culicoides halophilus	Kieffer, 1924	A	Morphological + PCR and sequencing
Culicoides helveticus	Callot, Kremer & Deduit, 1962	M	Morphological
Culicoides heteroclitus	Kremer & Callot, 1965	A, M, T	Morphological
Culicoides imicola	Kiefer, 1913	A, M, T	Morphological + PCR and sequencing
Culicoides indistinctus	Khalaf, 1961	T	Morphological
Culicoides jumineri	Callot & Kremer, 1969	A, M, T	Morphological + PCR and sequencing
Culicoides jurensis	Callot, Kremer & Deduit, 1962	A	Morphological
Culicoides kibunensis	Tokunaga, 1937	A, M, T	Morphological
Culicoides kingi	Austen, 1912	A, M, T	Morphological + PCR and sequencing + RFLP
Culicoides kurensis	Dzhafarov, 1960	A, M, T	Morphological
Culicoides landauae	Kremer, 1975	M	Morphological
Culicoides langeroni	Kiefer, 1921	A, M, T	Morphological + PCR and sequencing
Culicoides longipennis	Khalaf, 1957	A, M, T	Morphological
Culicoides marcleti	Callot, Kremer & Basset, 1968	A, M, T	Morphological
Culicoiodes maritimus	Kiefer, 1924	A, M, T	Morphological
Culicoides montanus	Shakirzjanova, 1962	A, M	Morphological + PCR and sequencing
Culicoides navaiae	Lane, 1983	A	Morphological
Culicoides newsteadi	Austen, 1921	A, M, T	Morphological + PCR and sequencing
Culicoides nubeculosus	Austen, 1921	A, M	Morphological
Culicoides nudipennis	Kiefer, 1922	A	Morphological
Culicoides. obsoletus	Meigen, 1818	A, M, T	Morphological + PCR and sequencing
Culicoides odiatus	Austen, 1921	A, M, T	Morphological
Culicoides oxystoma	Kieffer, 1910	M, T	Morphological + PCR and sequencing + RFLP
Culicoides pallidicornis	Kiefer, 1919	M	Morphological
Culicoides pallidus	Khalaf, 1957	M	Morphological
Culicoides paolae	Boorman, Mellor & Scaramozzino, 1996	A, M, T	Morphological + PCR and sequencing
Culicoides paradoxalis	Ramilo & Delécolle, 2013	A	Morphological
Culicoides parroti	Kiefer, 1922	A, M, T	Morphological
Culicoides pictipennis	Staeger, 1839	A, M	Morphological
Culicoides picturatus	Kremer & Deduit, 1961	A, M	Morphological
Culicoides poperinghensis	Goetghebuer, 1953	A	Morphological
Culicoides pseudojumineri	Salma, Chaker, and Babba, 2015	T	Morphological
Culicoides pseudolangeroni	Kremer, Chaker and Delécolle, 1981	T	Morphological
Culicoides pseudopallidus	Khalaf, 1961	A, M, T	Morphological
Culicoides pulicaris	Linnaeus, 1758	A, M	Morphological
Culicoides pumilus	Winnertz, 1852	M	Morphological
Culicoides punctatus	Meigen, 1804	A, M, T	Morphological
Culicoides puncticollis	Becker, 1903	A, M, T	Morphological + PCR and sequencing
Culicoides ravus	de Meillon, 1936	A, M	Morphological
Culicoides riethi	Kiefer, 1914	T	Morphological
Culicoides saevus	Kieffer, 1922	A, M, T	Morphological
Culicoides sahariensis	Kiefer, 1923	A, M, T	Morphological + PCR and sequencing
Culicoides santonicus	Callot, Kremer, Rault & Bach, 1966	A, M, T	Morphological
Culicoides schultzei	Enderlein, 1908	A, M	Morphological
Culicoides scoticus	Downes & Kettle, 1952	A, M	Morphological + PCR and sequencing
Culicoides sejfadinei	Dzhafarov, 1958	A, M, T	Morphological
Culicoides semimaculatus	Clastrier, 1958	A, M, T	Morphological
Culicoides sergenti	Kiefer, 1921	A, M	Morphological
Culicoides shaklawensis	Khalaf, 1957	A, M	Morphological + PCR and sequencing
Culicoides similis	Carter, Ingram & Macfe, 1920	M	Morphological
Culicoides simulator	Edwards, 1939	A, M	Morphological
Culicoides sp. (~C. enderleini)	Enderlein, 1908	A	Morphological + PCR and sequencing
Culicoides sphagnumensis	Williams, 1955	A	Morphological
Culicoides subfagineus	Delécolle & Ortega, 1998	M	Morphological + PCR and sequencing
Culicoides subfasciipennis	Kiefer, 1919	A, M, T	Morphological
Culicoides truncorum	Edwards, 1939	A, M	Morphological
Culicoides univittatus	Vimmer, 1932	A, M, T	Morphological
Culicoides vidourlensis	Callot, Kremer, Molet & Bach, 1968	M	Morphological
Forcipomyia alacris	Winn, 1852	A	Morphological
Forcipomyia bipunctata	Lineaus, 1767	A	Morphological
Forcipomyia biskraensis	Kieff, 1923	A	Morphological
Forcipomyia crassipes	Winn, 1852	A	Morphological
Forcipomyia formosae	Kieff, 1921	A	Morphological
Forcipomyia frutetorum	Winn, 1852	A	Morphological
Forcipomyia fuliginosa	Meigen, 1818	A	Morphological
Forcipomyia litoraurea	Ingram & Macfie, 1924	A	Morphological
Forcipomyia margaritae	Szadziewski, 1983	A	Morphological
Forcipomyia mesasiatica	Remm, 1980	A	Morphological
Forcipomyia monilicornis	Coquillett, 1905	A	Morphological
Forcipomyia murina	Winn, 1852	A	Morphological
Forcipomyia nigra	Winn, 1852	A	Morphological
Forcipomyia pallidipes	Santos Abreu, 1918	A	Morphological
Forcipomyia paludis	Macfie, 1936	M	Morphological
Forcipomyia phlebotomoides	Bangerter, 1933	A	Morphological
Forcipomyia picheyrei	Harant & Galan, 1942	A	Morphological
Forcipomyia pontica	Remm, 1968	A	Morphological
Forcipomyia psilonota	Kieffer, 1911	A	Morphological
Forcipomyia pulcherrima	Santos Abreu, 1918	A, T	Morphological
Forcipomyia rufescens	Kieffer, 1918	T	Morphological
Forcipomyia rugosa	Chan et Leroux, 1970	A	Morphological
Forcipomyia rustica	Kieff, 1919	A	Morphological
Forcipomyia sahariensis	Kieff, 1923	A	Morphological
Forcipomyia seneveti	Kieffer, 1922	A	Morphological
Forcipomyia striaticornis	Kieff, 1918	A	Morphological
Forcipomyia suberis	Clastrier, 1956	A	Morphological
Forcipomyia tenuisquama	Kieff, 1924	A	Morphological
Forcipomyia velox	Winn, 1852	A	Morphological
Forcipomyia waldemari	Szadziewski, 1983	A	Morphological
Forcipomyia wirthiana	Szadziewski, 1983	A	Morphological

A Algeria, M Morocco, T Tunisia.

.

While their diversity reflects their adaptability and ecological roles, biting midges’ economic impact is highly significant, mainly because of their ability to transmit pathogens, causing severe outbreaks. For instance, several Blue Tongue outbreaks have been reported in Algeria and Tunisia for several years, in 2000, 2006, and 2007, leading national veterinary authorities in the two countries to implement a series of control measures [84].

They are considered to be active transporters of several pathogens and well-known biological vectors of a number of arboviruses that cause over 100 worldwide human and veterinary infections (Figure 1b). So far, five pathogenic viruses have been detected in Ceratopogonids within the Maghreb region, including (i) the Epizootic Hemorrhagic Disease Virus (EHDV), two strains of which, EHDV-6 affecting cattle (B. taurus) and EHDV-8 affecting Barbary stag (Cervus elaphus barbarus), have been reported from Tunisia [85,86]. In contrast, reports from Algeria have not specified the strain associated with positive serology [87]. (ii) The African Horse Sickness Virus (AHSV), which has been detected in all three countries [88]. Additionally, the West Nile Fever Virus (WNFV), which classically affects both horses and humans, has been detected only in dromedary camels (Camelus dromedarius) in Tunisia and Morocco [89,90]. In Algeria, antibodies against the Akabane Virus (AKAV) have been detected in blood samples from dromedary camels [91].

It is impossible to mention the viral diseases transmitted by midges without alluding to the classical transmission of Blue Tongue Virus (BTV), as several studies conducted in the Maghreb have reported the detection of this virus in hosts, including small ruminants, cattle, and dromedary camels, as well as various species of Culicoides, including C. circumscriptus, C. imicola, C. newsteadi, and C. paolae. For instance, out of 21,175 susceptible sheep in Algeria, this disease resulted in approximately 2661 confirmed cases of clinically severe morbidity [92,93,94].

Moreover, Sghaier, Yaacoub [95] reported the presence of Mansonella perstans in non-resident students’ blood samples, raising questions about the possible dispersal of this blood parasite by biting midge species in North Africa.

The distribution map of medico-veterinary important midges in the Maghreb (Figure 2b) reflects their diverse taxonomic range. Culicoides circumscriptus and C. imicola are found across nearly all of the Maghreb, from the center of the Algerian Sahara and the Anti-Atlas region through the sub-Dahar mountains to the Mediterranean coastal areas of Morocco, Algeria, and Tunisia. In contrast, C. oxystoma is distributed from the southeastern Tunisian Atlas to the southeastern Aurès Mountains in Algeria and appears in the Moroccan Atlas’s western regions. C. newsteadi occupies the northeastern extremes of Algeria and northern Tunisia, spanning various ecoregions such as the Tell, the Aurès Massif, the Tunisian Atlas, and the western highlands of the Morocco and Tellian extensions. C. kingi shares a similar distribution pattern in Tunisia and Algeria with C. newsteadi, but extends further into the sub-Saharan areas in the south and the eastern coastal areas in northern Algeria. As for C. paolae, it is encountered in northern Tunisia and along the central Atlantic coast of Morocco. Although this species has not yet been reported in Algeria, it is believed that C. paolae may also inhabit northern parts of the country.

3.4. Tabanidae

Tabanidae, or “horseflies and deerflies”, are a large infraorder Tabanomorpha family, with approximately 4500 described species, with some genera being cosmopolitans [96]. Adults are characterized by stout bodies and large heads occupied mainly by compound eyes. The male’s holoptic eyes touch each other medially and take up the majority of the head, while, with the frons separating them, the female’s dichoptic eyes are smaller than the male’s.

In addition to the difference between males’ and females’ tabanids eyes, transverse stripes with yellow and green coloring patterns may be visible. The antenna comprises one broad basal segment and terminates in a canceled segment sequence [97]. So far, the explored literature census records mention 105 species of tabanids in the Maghreb listed in 10 genera. The females of most taxa are anautogenous, except for the Pangonius genus. Hence, 90 species of veterinary and medical importance are present within the considered geographic range (

Table 3. Checklist of morphologically identified tabanids with veterinary and medical importance in the Maghreb.

Genus	Species	Taxonomy	Country
Atylotus	Atylotus agricola	Wiedmann, 1828	M
	Atylotus agristis	Wiedemann, 1828	M, A
	Atylotus farinosus	Sziliidy, 1915	A
	Atylotus flavipes	Macquart, 1838	A
	Atylotus flavoguttatus	Sziliidy, 1915	A
	Atylotus fulvus	Meigen, 1820	M, A
	Atylotus kroberi	Surcouf, 1923	M, A
	Atylotus latistriatus	Brauer, 1880	M, A
	Atylotus loewianus	Villeneuve, 1920	M
	Atylotus pulchellus	Loew, 1858	A
	Atylotus quadrifarius	Loew, 1874	M, A
	Atylotus sublunaticornis	Zetterstedt, 1842	M
Chrysops	Chrysops caecutiens	Linnaeus, 1758	M
	Chrysops connexus	Loew, 1858	M
	Chrysops flavipes	Meigen, 1804	M, A
	Chrysops italicus	Meigen, 1804	M, A
	Chrysops mauritanicus	Costa, 1893	M, A
	Chrysops pallidiventris	Krober, 1922	M, A
	Chrysops relictus	Meigen, 1820	M
	Chrysops viduatus	Meigen, 1803	M
Dasyrhamphis	Dasyrhamphis barbata	Coscaron & Philip, 1967	M
	Dasyrhamphis algirus	Macquart, 1839	M, T, A
	Dasyrhamphis anthracinus	Meigen, 1820	M
	Dasyrhamphis ater	Rossi, 1790	M, A
	Dasyrhamphis carbonarius	Meigen, 1820	T
	Dasyrhamphis nigritus	Fabricius, 1794	M, A
	Dasyrhamphis tomentosus	Macquart, 1854	M, A
	Dasyrhamphis denticornis	Enderlein, 1925	A
	Dasyrhamphis goleanus	Szilady, 1923	A
	Dasyrhamphis villosus	Macquart, 1839	A
Ectinocerella	Ectinocerella. surcoufi	Seguy, 1925	M, A
Haematopota	Haematopota algira	Krober, 1922	M, A
	Haematopota benoisti	Séguy, 1930	M
	Haematopota bigoti	Gobert, 1881	M, A
	Haematopota crassicornis	Wahlberg, 1848	M
	Haematopota fusicornis	Becker, 1913	M
	Haematopota grandis	Macquart, 1834	M
	Haematopota italica	Meigen, 1804	M, A
	Haematopota lambi	Villeneuve, 1921	M
	Haematopota ocelligera	Krober, 1922	M, A
	Haematopota pandazisi	Krober, 1934	M, A
	Haematopota pluvialis	Linnaeus, 1761	M, A
	Haematopota subcylindrica	Pandellé, 1883	M
	Haematopota saccae	Leclercq, 1966	T
Heptatoma	Heptatoma pellucens	Fabricius, 1776	M
	Heptatomadecora	Loew, 1858	A (*)
	Heptatoma vittata	Fabricius, 1794	A
Hybomitra	Hybomitra arpadi	Szilády, 1923	M
	Hybomitra bimaculata	Macquart, 1826	M
	Hybomitra caparti	Leclercq, 1966	T
	Hybomitra distinguenda	Verrall, 1909	M
	Hybomitra macularis	Fabricius, 1794	M
	Hybomitra vittata	Fabricius, 1794	M
Silvius	Silvius algirus	Meigen, 1830	M, A
	Silviusalpinus	Scopoli, 1763	M
	Silvius variegatus	Fabricius, 1805	M
	Silvius appendiculatus	Macquart, 1846	A
Surcoufia	Surcoufia paradoxa	Krober, 1922	A, T
Tabanus	Tabanus albifrons	Szilacty, 1914	A
	Tabanus algirus	Meigen, 1830	A, T
	Tabanus algeriensis	Peus, 1980	A
	Tabanus autumnalis	Linnaeus, 1761	M, A
	Tabanus barbarus	Coquebert, 1804	M, A
	Tabanus auricpunctatus	Macquart, 1839	A
	Tabanus bifarius	Loew, 1858	M
	Tabanus bovinus	Linnaeus, 1758	M, A
	Tabanus bromius	Linnaeus, 1761	M, A
	Tabanus choumarae	Leclercq, 1967	M
	Tabanus crodiger	Meigen, 1820	M
	Tabanus cordigeroides	Surcouf, 1922	A
	Tabanus darimonti	Leclercq, 1964	M
	Tabanus eggeri	Schiner, 1868	M, A
	Tabanus dorsomaculatus	Macquart, 1847	A
	Tabanus guyonae	Surcouf, 1923	A
	Tabanus leleani	Austen, 1920	M, A
	Tabanus lunatus	Fabricius, 1794	M, A
	Tabanus maculicornis	Zetterstedt, 1842	M
	Tabanus mikii	Brauer, 1880	M
	Tabanus nemoralis	Meigen, 1820	M, A
	Tabanus mitidjensis	Macquart, 1838	A
	Tabanus quatuornotatus	Meigen, 1820	M
	Tabanus regularis	Jeannicke, 1866	M, A
	Tabanus rousseli	Macquart, 1839	M, A
	Tabanus seurati	Surcouf, 1922	A, T
	Tabanus spectabilis	Loew, 1858	M
	Tabanus spodopterus	Olsufjev, Moucha & Chvala, 1967	M
	Tabanus sudeticus	Zeller, 1867	M
	Tabanus unifasciatus	Loew, 1858	M
	Tabanus tinctus	Walker, 1850	M, A
	Tabanus albifacies	Loew, 1856	A

A Algeria, M Morocco, T Tunisia, (*) Unprecise mention of the taxon in the literature.

). The complete list, including species of marginal interest, can be consulted in the Supplementary Materials.

Tabanids’ mating occurs early in the morning during flight before the females seek a mammal host [98]. Females typically produce 100–800 eggs in one mass, and according to the species and the climate, embryogenesis can take from 2 to 21 days. However, eggs hatch quicker when the temperature and relative humidity are high [99]. Tabanus sp. female species need to take a blood meal after mating, which is necessary for egg maturation, except in autogenic species (e.g., Tabanus nigrovittatus) [100]. The lifecycle of tabanids is exclusively linked to their geographical distribution and climatic conditions. However, it generally starts with a single mass of a few hundred eggs that hatch in two or three weeks, laid by a female after a blood meal, but not necessarily in the case of non-hematophagous species such as Pangonius spp. or the case of autogenous species’ first oviposition [101,102]. Egg batches are often deposited on leaves or stems of vegetation in lotic, lentic, or even terrestrial habitats, contributing to the larvae’s morpho-ecological diversity. Tabanids larvae can be divided into the following three groups: rheophilic species with an affinity for rivers and streams, hydrobiont species inhabiting stagnant or slow-moving water bodies or mud, and edaphobiont species living in drier biomes such as forest litter [98,101].

The larval stage lasts up to three years for some species. Larvae exhibit different shapes and forms and every so often are predators of other larvae, worms, and small arthropods. Pupation occurs in dry places, lasting from one to three weeks and resulting in adults whose longevity is from about two to three weeks [98,101,102].

Tabanid flies have a high seasonality; they are more active in summer in temperate regions [103]. However, this activity varies considerably according to the biotope [104]. Such a phenomenon can be observed in Afrotropical regions, where tabanids are more active during December in the forest, while in the savannah, they are more observed in March [105].

Because of their hematophagy, various Tabanid species are suspected to be worldwide biological or mechanical vectors of several pathogens (Figure 1c), including viruses, bacteria, protozoans, and metazoans. They may also play a role in transmitting some Oestridae eggs. However, no studies on their vectorial role have been conducted so far in the Maghreb region [106,107].

Although not pathogenic, the biological and mechanical transmission of Trypanosoma theileri and T. kaiowa by Tabanids has been documented [108]. Given the high diversity and abundance of Tabanids in North Africa—acting primarily as mechanical vectors for trypanosomes—trypanosomiasis remains challenging to eradicate. Despite effective strategies targeting biological vectors and addressing trypanocidal resistance, non-integrated pest management practices continue to enable mechanical vectors to disseminate these pathogens, sustaining transmission in affected regions

Among the pathogens transmitted by tabanids (Figure 1c), the Equine Infectious Anemia Virus, “EIAV”, exclusively infecting equids and spreading at different pathogenesis stages through mechanical transmission, is transmitted by tabanids, such as Chrysops spp., Hybomitra spp., and Tabanus spp. [109,110]. Evidence of the mechanical transmission of African Swine Fever Virus (ASFV) by Tabanus hypomacros in China has also been reported [111]. Moreover, T. fuscicostatus has been proven to be a mechanical vector transmitting the bovine leukemia virus to small ruminants [112].

The lumpy skin disease virus, which causes a clinical state similar to foot-and-mouth disease (FMD), is often characterized by a high morbidity and low mortality among cattle. It can be mechanically transmitted by different species of blood-sucking dipterans, such as tabanids [113]. The virus was detected in various taxa of the Palearctic region and specifically on Haematopota spp in the Maghreb [51]. In August 2024, the United Nations Food and Agriculture Organization issued multiple warnings to North African and Euro-Mediterranean countries following the emergence of several outbreak sites in the southern, central, and eastern regions of northern Algeria. It is believed that this outbreak may have stemmed from the virus being introduced via Algeria’s southeastern borders, as a similar outbreak was reported in Libya in 2023 [114]. Other viruses, including Bovine Viral Diarrhea Virus (BVDV), have been reported as being naturally transmitted between cattle by T. fuscicostatus [115].

The filarial Loa loa, known as the tropical eye worm and an agent of human loiasis, is an endemic parasite limited to the equatorial rainforests of Africa. It is biologically transmitted by Chrysops spp., commonly known as deer flies. Chrysops silacea and Chrysops dimidiata are the main vectors of human loiasis, while other Chrysops species are mainly zoophilic. In the Republic of Congo, C. dimidiata is the principal vector in the forest, whereas C. silacea is the major vector in cleared forest zones near villages [101]. Moreover, Tabanus and Hybomitra species are considered to be intermediate hosts for Elaeophora schneideri, a filarial nematode of domestic and wild ruminants in North America, occurring commonly in mule deer, elk, and moose, as third-stage larvae have been observed in adults [116,117].

Some neotropical bot flies, such as Dermatobia hominis (Diptera: Oestridae)—commonly known as the “Torsalo” in Mexico, Argentina, and Brazil, or the “American warble fly”—are responsible for causing furuncular myiasis in humans and animals. These flies use a unique strategy for egg transport, where they attach their eggs to various arthropods, including Tabanid flies, which then carry the eggs to potential infection sites [118,119]. The vectorial role of Tabanids extends beyond transmitting parasitic worms; they are also efficient vectors for various bacterial species. For instance, Francisella tularensis, the bacterium responsible for tularemia, is transmitted by multiple arthropod vectors, including Tabanid species from the Tabanus and Chrysops genera and, more specifically, C. discalis [120].

Conversely, Bacillus anthracis, the causative agent of anthrax, can be transmitted through direct contact with dead animals harboring spores or mechanically by Tabanids to various mammals, including humans. This was illustrated in a case report by Fasanella, Di Taranto [121] in Italy, where a breeder contracted anthrax after being bitten by Tabanids, despite having had no direct contact with dead animals. Another example of a bacterial disease of veterinary importance transmitted by Tabanids is anaplasmosis due to Anaplasma marginale, a rickettsial pathogen primarily transmitted biologically by ticks. Nevertheless, horseflies are considered to be mechanical vectors for this pathogen and are believed to play a role in its transmission to livestock [122]. Other bacteria, such as Brucella spp., Pasteurella multocida, Erysipelothryx rhusiopathiae, or Ehrlichia risticii, have also been associated with tabanids in experimental transmission [123,124].

Regarding protozoan transmission, Besnoitia besnoiti can survive for up to 24 h in the mouthparts of certain African Tabanid species, such as Atylotus nigromaculatus and Tabanocella denticornis, facilitating its mechanical transmission. Regarding Trypanosoma species, Tabanids can act as both mechanical and biological vectors, with the mode of transmission depending on the specific Tabanid and Trypanosoma species involved in sontigun [124].

Trypanosoma vivax, the causative agent of Nagana in cattle and wild mammals, has been experimentally shown to be mechanically transmitted by Afrotropical Tabanid species, including Atylotus agrestis and A. fuscipes [125,126]. Additionally, the DNA of Trypanosoma congolense, another Nagana-causing agent in domestic animals, was recently identified in Tabanids in Mozambique [127]. Meanwhile, Trypanosoma evansi, the agent of Surra—which primarily affects camelids and equids and has spread from North Africa to the Middle East, Asia, and South America through the movement of infected animals [128]—can also be mechanically transmitted by Tabanids, including Atylotus agrestis, Dasyrhamphis tomentosus, and Tabanus nemoralis [41,123,129]. This transmission was the major route for the spread of the disease in India, where the high prevalence of Surra was associated with an increase in the population of tabanids [126].

Tabanids’ distributional profiles are as varied as their taxonomic record, behavior, hosts, and environment-dependent factors. In Morocco, the range of horseflies and deerflies primarily extends along the central high plain to the Anti–Atlas axis. However, species from the Hybomitra and Chrysops genera are restricted mainly to the southern part of the High Atlas, with Hybomitra species also occurring at lower altitudes and in coastal Atlantic areas. In the eastern Maghreb, the genus Tabanus has the widest range, spanning from Algeria’s central coastal areas through the Tunisian Atlas, the Aurès Massif, and into northern and southeastern Tunisia’s coastal regions. This range even reaches the northeastern Sahara, where the genus Haematopota can be observed. This genus is also present along the entire eastern Algerian coast. Conversely, species within the Dasyrhamphis genus occupy much of northeastern Algeria and parts of northwestern Tunisia. Meanwhile, the ranges of the Chrysops and Atylotus genera overlap considerably, confined to northeastern Algeria’s humid and sub-humid coastal regions (Figure 2c).

3.5. Phlebotominae

Phlebotominae (Diptera: Psychodidae), commonly known as “Sandflies” in different world regions, represent a monophyletic group of hematophagous dipterans. This subfamily includes approximately 900 taxa, about 70 of which have been reported as vectors of leishmaniasis [130]. In the Maghreb (Algeria, Morocco, and Tunisia), 29 species have been progressively identified over a century (Supplementary Materials). These latter are affiliated with one of two genera, Phlebotomus and Sergentomyia, and cover 15 and 14 species within these countries (

Table 4. Checklist of phlebotominae in the Maghreb with their respective identification methods.

Genus	Taxon	Taxonomy	Country	Identification
Phlebotomus	Phlebotomus bergeroti	(Parrot, 1934)	M, A, T	M
	Phlebotomus papatasi	(Scopoli, 1786)	M, A, T	M + MT + seq
	Phlebotomus ariasi	(Tonnoir, 1921)	M, A, T	M
	Phlebotomus chadlii rioux	(Juminer et Gibily, 1966)	M, A, T	M
	Phlebotomus langeroni	(Nitzulescu, 1930)	M, A, T	M
	Phlebotomus longicuspis	(Nitzulescu, 1930)	M, A, T	M + MT + seq
	Phlebotomus mariae	(Rioux, Croset, Léger et BaillyChoumara, 1974)	M	M
	Phlebotomus perfiliewi s.l.	(Parrot, 1930)	M, A, T	M + MT + seq
	Phlebotomus perniciosus	(Newstead, 1911)	M, A, T	M + MT + seq
	Phlebotomus alexandri	(Sinton, 1928)	M, A, T	M
	Phlebotomus chabaudi	(Croset, Abonnenc et Rioux, 1970)	M, A, T	M
	Phlebotomus kazeruni	(Theodor et Mesghali, 1964)	M, A	M
	Phlebotomus riouxi	(Depaquit, Léger et Killick-Kendrick, 1998)	M, A, T	M
	Phlebotomus mascittii	(Grassi, 1908)	A	M
	Phlebotomus sergenti	(Parrot, 1917)	M, A, T	M + MT + seq
Sergentomyia	Sergentomyia antennata	(Newstead, 1912)	M, A, T	M
	Sergentomyia bedfordi	(Newstead, 1914)	M	M
	Sergentomyia fallax	(Parrot, 1921)	M, A, T	M + MT + seq
	Sergentomyia minuta	(Rondani, 1843)	M, A, T	M + MT + seq
	Sergentomyia schwetzi	(Adler, Theodor et Parrot, 1929)	M, A, T	M
	Sergentomyia christophersi	(Sinton, 1927)	M, A, T	M
	Sergentomyia clydei	(Sinton, 1928)	M, A, T	M
	Sergentomyia a. asiatica	(Newstead, 1912)	M	M
	Sergentomyia lewisi	(Parrot, 1948)	M, A, T	M
	Sergentomyia dreyfussi	(Parrot, 1933)	M, A, T	M
	Sergentomyia a. eremitis	(Parrot et de Jolinière, 1945)	A	M
	Sergentomyia hirtus	(Parrot et de Jolinière, 1945)	A	M
	Sergentomyia tiberiadis	(Adler, Theodor et Louric, 1930)	A	M
	Sergentomyia cincta	(Parrot and Martin 1948)	A	M

A Algeria, M Morocco, T Tunisia. Identification tool: M Morphological, MT Maldi Tof, Seq Sequencing.

).

Phlebotominaes’ lifecycle is holometabolous, lasting 30 days on average [131]. After laying eggs, they hatch within 1–2 weeks, and the terrestrial saprophage larvae, during all four instars, colonize moist soil with an affinity for cool temperatures. Pupation starts after three weeks. The formed pupa hatches within 10 days, producing adults ready to mate. Palearctic sandfly species exhibit unique adaptations, such as the ability to overwinter by entering diapause in the final larval instar, allowing them to survive unfavorable conditions [131,132].

In addition to their diversity, shaping, in part the panoply of their seasonality and biology, sandflies are very sensitive to abiotic factors that impact their charge in the environment, including those related to human agricultural activity such as irrigation and animal breeding, as well as landscape and meteorological factors [133,134]. For instance, in the Mediterranean subregions, sandflies can be present in several natural microhabitats, exhibiting a characteristic seasonal activity with a peak in dry seasons [135]. Their activity is mainly nocturnal (from 9 p.m. to 1 a.m.) in indoor areas with a low barometric pressure [136].

The feeding behavior of phlebotominae differs by sex, as females are hematophagous, while males are phytophages. Phlebotomine sandflies can be identified through morphological techniques, focusing on male genital structures and female spermathecae. However, morphometric methods are ineffective for distinguishing species within the Phlebotomus perniciosus (Newstead, 1911) complex. Consequently, innovative molecular techniques, including MALDI-TOF, DNA barcoding, and eggshell ultrastructure, are becoming a trend in sandfly taxonomy [137,138]. Overall, these approaches might serve to broaden the knowledge of phlebotome diversity.

Because their biology is closely tied to environmental and human activities, sandflies play a critical role as vectors of serious diseases such as leishmaniasis. Their vectorial role in transmitting leishmaniasis is not documented in all taxa found within the Maghreb region [139]. Cutaneous leishmaniasis (CL), the predominant form of this disease, caused by Leishmania major and L. tropica, is transmitted exclusively by Phlebotomus papatasi (Scopoli, 1786) and Ph. sergenti (Parrot, 1917). In contrast, Ph. perniciosus, Ph. ariasi (Tonnoir, 1921), and Ph. longicuspis (Nitzulescu, 1930) are recognized as vectors of L. infantum, known to cause both visceral leishmaniasis (VL) and CL [139,140]. However, L. infantum was also detected in Ph. langeroni (Nitzulescu, 1930), Ph. longicuspis, Ph. papatasi, Ph. perfiliewi (Parrot, 1930), and Sergentomyia minuta (Rondani, 1843) [141,142].

The cited leishmaniasis agents mainly require small vertebrate reservoirs for their persistence in the environment. Several studies from Algeria, Morocco, and Tunisia have reported the natural infection of rodents, including Rattus norvegicus, R. rattus, Mus musculus, Meriones shawi, Psamomys obesus, Massoutiera mzabi, and Ctenodactylus gundi [143,144]. However, this list remains open, as studies have identified other wild rodents, such as Gerbillus amoenus, as potential competent hosts for leishmanias [145]. Moreover, other micromammals, such as hedgehogs (Atlerix algirus and Paraechinus aethiopicus), have been reported as natural reservoirs for L. major [146].

As for phleboviruses, nine genotypes have been detected in sandflies sampled from the Maghreb, which are, to date, as follows: Medjerda valley virus (MVV), Punique virus, Saddaguia virus (SADV), Sandfly fever Naples virus (SFNV), Sandfly Fever Sicilian Virus (SFSV), Sandfly fever turkey Virus (SFTSV), Utique virus (UTIV), Uukuniemi phlebovirus (UUKV), and Toscana virus (TOSV) (Supplementary Materials). The phleboviruses detected in the Maghreb and other Mediterranean European countries infect most humans and dogs and have also been detected individually or in pools of the following sandflies: Ph. ariasi, Ph. perfiliewi, Ph. perniciosus, Ph. longicuspis, Ph. langeroni, Ph. papatasi, and Se. minuta. Figure 1d represents all the associations of the sandflies with their hosts and related pathogens reported in the three countries.

The six important sandfly vectors encountered in the Maghreb exhibit distinct distribution profiles. Ph. perniciosus has the broadest range, covering nearly all Moroccan and Tunisian territories and extending across much of the eastern Sahara. Ph. longicuspis and Ph. sergenti share a similar range, spanning from the Atlas Belt and expanding into the northeastern part of Algeria’s Sahara, thus covering various ecoregions (humid, sub-humid, semi-arid, arid, and desertic areas). Similarly to Ph. Perniciosus, Ph. perfiliewi appears to be the only species confined exclusively to humid and sub-humid regions, from north-western Algeria to northeastern Tunisia. Finally, Ph. langeroni and Se. antennata exhibit very similar distribution profiles, being found in the northern High Atlas in Morocco, the eastern Sahara, the Aures and Tunisian Atlas, and the east coast of the Maghreb (Figure 2d).

3.6. Simuliidae

Simuliidae, or “Black flies”, are medically and veterinary relevant invertebrates that depend on benthic streams [147]. This family includes 2408 living species organized into 31 genera [25,148]. Within this family of dipterans, taxa with similar morphological characteristics can lead to misidentification, which limits the understanding of their ecology and vectorial roles [149]. Indeed, the classification of blackflies often requires a holistic approach considering different characters of immature stages, adult males, females, giant polytene chromosomes (wide and large chromosomes found in some dipteran larvae, usually n = 3), and DNA barcoding, as well as ecological data [150,151].

Studying the patterns of giant chromosomes, often developed in the silk gland cells of the larval stages, reveals that many taxa regarded as distinct species are complexes of different biologically unique entities known as sibling or cryptic species, which refers to genetically distinct and morphologically similar organisms, making simulids the most genetically characterized living group [152]. However, this observation leverages the need for more research on the vectorial competency of the various species of simuliids.

Another aspect of peculiar interest in studying simuliids is their holometabolous lifecycle. Mating flights and crawling at low heights during reproduction are common behaviors observed in various species of black flies [150,153,154]. These behaviors may be influenced by factors such as the small size of compound eyes and the absence of phototaxis in certain species, which could contribute to the lack of mating flights in these populations. Additionally, some species exhibit parthenogenesis (an asexual reproduction in which the embryo develops directly from an egg without the need for fertilization) either exclusively or alongside mating, leading to the observed imbalances in the male-to-female sex ratio [155,156]. Despite mating attempts, females frequently reject males during the copulation process, which may hamper successful reproduction. Moreover, external fertilization is also possible, as males have been observed thrusting and crawling on laid eggs using their claspers [150,156].

Once ready, females lay thousands of oval- or triangular-shaped eggs, except for some species producing approximately one billion eggs per kilometer of riverbed daily. This high number of individuals within a specific region makes blackflies one of the few arthropods that have killed animals via exsanguination and toxemia during massive attacks [150]. After hatching, larvae attach by their posterior proleg to a silk pad applied to substrates on river rocks and extend their body to filter feed from water. Silk larvae are secreted by several glands at almost all lengths of their bodies [157]. Some studies have correlated the adhesiveness of different species’ silk with the velocity of flowing water [158]. During feeding, larvae filter nutrients with the anterior labral fans attached to their sclerotized head. This filter-feeding habit and their respiratory needs are probably some drivers of the various distribution profiles of different taxa [150].

It is worth noting that immature stages generally occur in aquatic environments. However, the pupa of many species have terrestrial adaptations, making them resistant for years if the water recedes [150].

In North African countries (Algeria, Morocco, and Tunisia), six genera are known so far, which are, to date, Greniera, Helodon, Metacnephia, Prosimulium, Simulium, and Urosimulium (

Table 5. Checklist of important blackflies reported in the Maghreb.

Genus	Taxon	Taxonomy	Country	Identification
Helodon	Helodon laamii	Beaucournu-Saguez & Bailly-Choumara	M	M
Metacnephia	Metacnephia blanci	(Grenier & Théodorides, 1953)	M, A, T	M
Prosimulium	Prosimulium aculeatum	(Grenier & Bailly-Choumara, 1972)	M	M
	Prosimulium albense	(Rivosecchi, 1961)	A	M
	Prosimulium fungiforme	(Adler, Belqat, Haouchine, 2024)	A	M + CC
	Prosimulium hirtipes	(Beaucournu-Saguez & Bailly-Choumara, 1981)	M	M
	Prosimulium latimucro	(Enderlein, 1925)	M	M
	Prosimulium rufipes	(Meigen, 1830)	M, A (*)	M + CC
	Prosimulium tomosvaryi	(Enderlein, 1921)	M	M + CC
Simulium	Simulium angustipes	(Edwards, 1915)	M, A, T	M
	Simulium angustitarse	(Lundström, 1911)	M, A	M
	Simulium argenteostriatum	(Strobl, 1898)	A, T	M
	Simulium atlasicum	(Giudicelli & Bouzidi,1989)	M	M
	Simulium auricoma	(Meigen, 1818)	M	M
	Simulium aureum	(Fries, 1824)	A	M
	Simulium berberum	(Giudicelli & Bouzidi 1989)	M	M
	Simulium bezzi	(Corti, 1914)	M, A	M
	Simulium brevidens	(Rubzov, 1956)	M, A	M
	Simulium carthusiense	(Grenier & Dorier, 1959)	M	M
	Simulium costatum	(Friederichs, 1920)	M, A	M
	Simulium cryophilum	(Rubzov, 1959)	M, A, T	M + CC
	Simulium egregium	(Séguy, 1930)	M	M
	Simulium erythrocephalum	(De Geer, 1776)	T	M
	Simulium equinum	(Linnaeus, 1758)	M, A (*)	M
	Simulium galloprovinciale	(Giudicelli, 1963)	M, A	M
	Simulium gracilipes	(Edwards, 1921)	M, A	M
	Simulium hispaniola	(Grenier & Bertrand, 1954)	A	M
	Simulium ibleum	(Rivosecchi, 1966)	M, A, T	M
	Simulium intermedium	(Roubaud, 1906)	M, A, T	M
	Simulium knidirii	(Doby & David, 1960)	M	M
	Simulium lamachei	(Doby & David, 1960)	M	M
	Simulium latipes	(Meigen, 1804)	T (*)	M
	Simulium lundstromi	(Enderlein, 1921)	M, A	M
	Simulium marocanum	(Bouzidi & Giudicelli, 1987)	M, A	M
	Simulium mellah	(Santos Abreu, 1922)	M, A	M
	Simulium monticola	(Friederichs, 1920)	A	M
	Simulium ornatum	(Meigen, 1818)	M, A, T	M
	Simulium petricolum	(Rivosecchi, 1953)	M, A	M
	Simulium pseudequinum	(Séguy, 1921)	M, A, T	M
	Simulium quadrifila	(Grenier, Faure & Laurent, 1957)	M, A	M
	Simulium reptans	(Linnaeus, 1758)	T	M
	Simulium rubzovianum	(Sherban, 1961)	A, T	M
	Simulium ruficorne	(Macquart, 1838)	M, A, T	M
	Simulium sergenti	(Edwards, 1923)	M, A, T	M
	Simulium toubkal	(Bouzidi & Giudicelli, 1986)	M	M
	Simulium trifasciatum	(Curtis, 1839)	M, A	M
	Simulium variegatum	(Meigen, 1818)	M, A	M
	Simulium velutinum	(Santos Abreu, 1922)	M, A	M
	Simulium vernum	(Macquart, 1826)	M, T	M + CC
	Simulium weiningense	(Chen & Zhang, 1997)	M, A	M
	Simulium xanthinum	(Edwards, 1933)	M, A	M
Urosimulium	Urosimulium faurei	(Bernard, Grenier & Bailly-Choumara, 1972)	M, A	M
	Urosimulium jucci	(Contini, 1966)	A (*), T	M
Greniera	Greniera fabri	(Doby & David, 1959)	A	M

M morphology, CC Cytogenic confirmation, (*) Unique unprecise mention of the taxon in literature.

). The most important genus from an epidemiologic perspective is Simulium, not only because of the high number of species belonging to it (n = 1979), but because over 90% of vector simuliids are within this genus [25]. However, most Maghrebin species are not incriminated in the transmission of diseases. For instance, 54 species have been recorded in the Maghreb (42 in Morocco, 37 in Algeria, and 16 in Tunisia). Some of them, such as Simulium brevidens, S. cryophilum, and S. vernum, belong to the S. vernum species group, in which many members have been reported as vectors of various protozoans, including Leucocytozoon cambournaci, L. dubreuili, L. lovati, L. toddi, L. ziemanni, and Trypanosoma avium, which infect, respectively, sparrows, thrushes (Turdus spp.), grouse (Turdus Philomelos and Lyrurus tetrix), imperial eagles (Aquila heliacal), hawks (Accipiter gentilis), owls (Strigidae), and buzzards (Buteo buteo) in different parts of the world of the Palearctic region (Supplementary Materials).

Avian protozoans have also been associated with the Simulium aureum (Fries, 1824) complex, with several cytospecies identified in the Maghreb (S. angustipes (Edwards, 1915), S. erythrocephalum (De Geer, 1776), S. petricolum (Rivosecchi, 1953), S. rubzovianum (Sherban, 1961), S. velutinum (Santos Abreu, 1922), and S. weinengense (Chen & Zhang, 1997)). These pathogens, including L. cambournaci, L. dubreuili, L. icteris, L. lovati, L. sakharoffi, L. smithi, L. toddi, L. ziemanni, and T. avium, specifically infest various bird species such as sparrows, thrushes, blackbirds, grouse, corvids, turkeys, hawks, owls, raptors, chickens, and guinea fowl [150].

Notwithstanding L. caulleryi, transmitted by biting midges (Ceratopongidae), all leucocytozoonosis agents are transmitted in avian populations by black flies, mainly through bites.

The haemosporidian reported in the upper western and extreme eastern Palearctic countries (Supplementary Materials) may have a wider geographical distribution due to the cosmopolitan circulation of many bird species during nesting or hibernating seasons. Since most leucocytozoons infest wild birds for their whole life, the host generalist behavior observed by ornithophilic simuliids also contributes to the dispersal of leucocytozoons in different parts of the world. These observations enhance our hypothesis about the circulation of different leucocytozoons in the Maghreb region, yet a molecular survey is necessary to confirm this hypothesis.

Although most pathogens transmitted by blackflies (Figure 1e) primarily infest avian species, others, such as Onchocerca lienalis and O. volvolus, infesting cattle and humans, respectively, are considered to be a major concern in Africa. Notably, O. volvulus, which is the second leading cause of infectious blindness worldwide, is exclusively transmitted by Central African black fly species such as S. damnosum, a vector species that is not encountered in the Maghreb region [25,150]. However, S. erythrocephalum, one of the vectors of O. lienalis infesting cattle, has been reported in Tunisia [159], underscoring the need to study simuliid-borne pathogens in the Maghreb.

Human onchocerciases are a severe public health and economic issue, despite their low incidence. Several cases, including O. lupi infecting humans, diagnosed in the Maghreb region, were previously reported in Tunisia [160].

Notwithstanding the high diversity of simuliids in the Maghreb, which may play a crucial role in maintaining the disease in the environment, other attributes such as altitude, temperature, anautogeny, and repetitive gonotrophic cycles in some species characterized by multi-voltinism and anthropophily, as well as the ability of some species to colonize large rivers, set the scene for black fly distribution and their borne diseases circulation [150,155]. The distribution of medically and veterinary important blackflies is heterogeneous, with certain species reported in some regions, but not others. Notably, S. pseudequinum is found in the western part of the Maghreb, encompassing the entire Mediterranean coast, high plains, Middle and High Atlas, and Anti–Atlas regions of Morocco. On the other hand, S. velutinum, found in humid and sub-humid regions, covers the entire Mediterranean coast of Morocco and the northeastern Algerian coast, Djurdjura, Aurès, and the high plains. Notably, S. aureum has a similar range to S. velutinum, but has only been reported in Algeria. As for S. rubzovianum, it is restricted to the northern humid regions of Tunisia and the far eastern regions of Algeria (Figure 2e).

4. Conclusions

In the Maghreb, up to October 2024, 324 taxa of biting flies and midges have been reported, some of which are endemic to the region, while others can be encountered in the Palearctic and Afrotropical regions. This study emphasizes these insects’ ecological complexity and notable diversity, especially their role as vectors of pathogens with medical and veterinary relevance. The most significant groups from an epidemiological perspective include Phlebotominae, associated with leishmaniasis, and Ceratopogonidae, involved in viral diseases such as the Blue Tongue virus. However, other taxa, including blackflies, tabanids, and stable flies, remain less studied in this region, underscoring critical gaps in epidemiological knowledge. From deserts to humid Mediterranean zones, the Maghreb’s distinct ecological and climatic diversity is reflected in the geographic distribution of these insects. Across several regions, Phlebotomus perniciosus and Culicoides imicola illustrate the risk of disease transmission in changing environmental and climatic conditions.

Additionally, this study highlights the need for integrative techniques that combine genomic and morphological approaches to improve vector surveillance and taxonomic resolution. Nevertheless, more investigation is required to understand the epidemiological relevance and vectorial roles of poorly defined species, such as Simuliidae and Tabanidae. Region-specific integrated vector management strategies must be established to reduce disease risks and improve public and veterinary health in the Maghreb. These strategies must include monitoring emerging pathogens and their interactions with hosts.

The present study could guide entomologists and epidemiologists interested in biting flies and their control by providing a foundation for future research and policy making by mapping their distribution and identifying knowledge gaps. It comprehensively explains the region’s biting fly biodiversity and health and ecosystem management implications.