Ultrastructural and molecular approach as a tool for taxonomic identification of aquatic macroinvertebrates: A review

Abstract

Aquatic insects require water at one or other phase for the completion of their life cycle. The insect larvae serve as food for larger invertebrates and vertebrates in aquatic food chain. Their diversity, number, and abundance act as water quality indicators, and thus species are classified accordingly as pollution tolerant or sensitive. So, identifying these aquatic larvae and macroinvertebrates are important for determining the biodiversity, and classification of insect species, followed by assessment of water health, and understanding the influence of climate change and anthropogenetic activities on these. Chances of misidentification have been reported due to loss of expertise, absence of taxonomic keys for larvae or intermediate stages, or damaged structure during collection or preservation. Recent advances in molecular and electron microscopy have revolutionized the identification procedure. Scanning electron microscopy detail the structure and morphology of the insect, while molecular techniques employing PCRs, DNA barcoding, and molecular markers allow the identification of the insects from any tissue (whole/part), and comparing the nucleotide sequences helps in the evaluating the family tree and lineage. The review summarizes the present status of aquatic invertebrates identification and the importance of these two techniques in the taxonomic identification of aquatic insects.

Aquatic insects; Macroinvertebrates; Molecular markers; Electron microscopy; DNA barcoding.

1. Introduction

Aquatic invertebrates form an important component of the freshwater ecosystem. It comprises organisms lacking backbones varying from μm to cm in size and has a great diversity of immature and adult aquatic fauna. Freshwater bodies contain a large variety of macroinvertebrates from various phyla such as arthropods (insects, scuds, and crayfish), annelids (worms and leeches), mollusks (snails, clams, and mussels), and nematodes (roundworms) (Hauer and Resh, 2017). These are generally larger than 2 mm in size (captured through 500mm of mesh size) and exhibit a pivotal role in the aquatic food chain and energy flow in the system (Wallace and Webster, 1996; Beermann et al., 2018a). These may be benthic i.e., linked to channel bottom surfaces (rock, cobble, and thinner sediments) or other fixed surfaces (collapsed trees, fruit trees, roots, submerged and emergent aquatic vegetation) (De Pauw et al., 2006). Regular monitoring and bio-assessment are tools for ecosystem management and determination of the biological integrity of freshwater streams (Sharma and Chowdhary, 2011). A combination of physical, chemical, and biological parameters determine stream health (Reynoldson et al., 1989; Verneaux et al., 2003). In the biological method, macroinvertebrates make the temporal scale of the appropriate response related to anthropogenic activities. These are preferred over zooplankton and phytoplankton due to their relatively long life cycle (Rosenberg, 1993).

Valuable features of macroinvertebrate communities include enormous diversity, low mobility, and longer life spans; ability to show responses towards numerous types and levels of pollutants; non-hassle collection and identification up to family level; and are an important link in food webs (Pavluk et al., 2000; Bonada et al., 2006; Morse et al., 2007). So, the determination of species diversity, biotic indices, and functional feeding groups act for biomonitoring of lotic waters (Li et al., 2010; Tomanova et al., 2007). Macro-invertebrates can tolerate varied aquatic conditions and pollution levels that mark their suitability as indicators (Morse et al., 2007; Pavluk et al., 2000). Due to the small size and varied morphologies (larval forms) of macroinvertebrates, there arises a need to identify them correctly (accurately) which requires the employment of advanced techniques such as Electron microscopy and molecular techniques. The present review aims at summarizing the current status of aquatic invertebrates identification and the importance of these advanced techniques in the taxonomic identification of aquatic insects.

Methodology: For the present article, works of literature (research, review, and books), pertaining to macroinvertebrates, aquatic organisms, electron microscopy, and molecular techniques were searched on different databases like google scholar, research gate, and scopus. Finally compiled the article under various headings and subheadings.

2. Impact of physiological and chemical parameters of water on macroinvertebrates

The ecology of freshwater depends on various physiological and chemical parameters like pH, temperature, dissolved oxygen (DO), water current, and conductivity (Jindal and Singh, 2020). Temperature is among the most critical environmental factors impacting benthic macroinvertebrates distribution and community structure (Sharma et al., 2015). In the case of aquatic insects, the emergence pattern, growth rate, metabolism, breeding, and body size are equally affected (Tali et al., 2013). Many macroinvertebrate species show variation in tolerating temperature ranges (Angelier, 2003). However, global warming is affecting the communities and causing the loss of cold-water species belonging to the order Ephemeroptera (Mayflies), Plecoptera (stoneflies), and Trichoptera (caddisflies) in the USA, Atlantic, and Korea, which may lead to the loss of food in the system, affect nutrient recycling and may even result in the replacement of the community by invasive one (Hamilton, 2010).

Water current is a significant characteristic of running water. In terms of ecology, morphology, and behavior, stream creatures vary from their still-water counterparts in their capacity to adapt to continually moving water rather than those living in stagnant waters (Giller et al., 1998; Crisci-Bispo et al., 2007). Many invertebrate species have been wiped away by water currents due to their inability to withstand high velocity (Ramírez and Pringle, 2001). While during low water flow time macroinvertebrates may move into areas of faster streamflow (Suren and Jowett, 2006). For example, with an increase in current flow, the percentage of chironomid larvae dropped among the benthos of Hawaiian streams (Kinzie et al., 2006).

Dissolved gases such as oxygen and carbon dioxide are crucial constituents of the freshwater system and are controlled by other factors like temperature, partial pressure of gases, respiration, salinity, and photosynthesis (Allan 1995; Wetzel and Likens 2000). Benthic macroinvertebrate species can tolerate varied concentrations of dissolved oxygen (DO) (Connolly et al., 2004; Sharma and Rawat, 2009). In winter, associated with low temperatures, raised values of DO and higher populations of macroinvertebrates have been recorded (Negi and Mamgain, 2013). The pH of the water stream significantly varies between 5.0 - 9.0 and directly influenced by human activity and environmental conditions (Hussain, 2012). Alterations in pH directly impact the population (Yuan, 2004). Moreover, the productivity of the aquatic freshwater habitats has substantially declined at lower pH (5.0), thereby reducing food availability to higher organisms (Thomsen and Friberg, 2002). Most macroinvertebrates had explicated detrimental effects at pH levels above 10 at one or more stages of life.

Conductivity measures the concentration of charged ions present in the water and is greatly affected by the circumstances of the terrain. Conductivity has been identified as one of the major elements influencing the distribution and composition of mayflies and stoneflies (Hrovat et al., 2009). Species richness declines with an increase in stream conductivity (Pond et al., 2008). In the study by Wallace and Eggert (2009) the abundance of the vulnerable benthic species has been reported to be reduced or replaced with pollution-tolerant dipterans (such as Trichoptera, Ephemeroptera, and Plecoptera) associated with conditions of increased conductivity.

3. Identification of macroinvertebrates

Traditionally, the identification of a species (organism) involves identifying and comparing morphological characters using taxonomic keys. Although morphological characteristics remain frequently used in tracking biodiversity, it presents certain disadvantages (Baird and Hajibabaei, 2012). Morphological keys are based on recognizable traits of the adult but at times cannot distinguish larval phases or early stages of development as observed in the case of aquatic insects. It is either due to the non-availability of keys or sample damage incurred during sorting (Sweeney et al., 2011). For taxonomic identification through keys, there is always a need to consult the taxonomist to certify the accurate assignment of species. Inadvertently there may incur misidentification of the species up to 7.5–9%, even if performed by experienced taxonomists (Metzeling et al., 2002; Hewlett, 2000; Sweeney et al., 2011). To overcome the shortcomings advanced morphological identification techniques such as SEM (Scanning Electron Microscopy) and molecular characterization can be employed (Hübner et al., 2017).

4. Ultrastructural identification of macroinvertebrates

The use of scanning electron microscopy in biological studies has a history dating back 1970s (Kownachi et al., 2015). SEM has been employed to image the surface morphology, topography, and composition of non-biological and biological materials with maximum magnification up to 5,00,000X and spatial resolution below 1nm (Grover et al., 2022; Wang, 2000; Wang et al., 2000). It examines the whole specimen under an electron microscope and characterizes microscopic details that aid in identifying the species (Figure 1). It has developed an understanding of the function of various structures (Ortiz et al., 2015). Imaging through SEM has helped in the detailed characterization of morphological features such as the structure of hooklets, holes on reticular structures, the plastron plate of the thoracic horn, and ejaculates in Chironomids (Kownacki et al., 2015) (Table 1). It has been used to differentiate two closely related species, as evidenced by an SEM micrograph of the claws of two different species (Fig. 2a,b) (Jindal and Singh, 2020).

Figure 1.

Procedure for SEM sample preparation.

Table 1.

Identification and characterization of macroinvertebrates through SEM.

Study site	Morphological characters	Invertebrates	Significance	References
Binwa Stream India	•Serrations on inner side of tarsal claws •Presence of sensilla in proximity to the upper surface of tarsal claw •suckers and serrated setae on ventral side near the external gills in B. Himalayana •Adhesive apparatus (first time reported)	Baetis bifurcates Baetis himalayana	•Differentiated two similar species •Adhesive apparatus (first time reported)	Jindal and Singh (2020b)
	•Presence of paired (bifid) tarsal claw and proleg claw in the species	Cryptoperla sp.	Characterized morphological characters in detail
	•Possess clamp like structure on the tarsal claw	Rhyacophila sp.,
	•Thin, long, paired mesoleg and metaleg tarsus	Naucoris sp.
	•Friction pads on the lateral side, with numerous serrated setae	Ecdyonurus sp.
Krakow, Poland	•presence of holes on the reticular structure •peculiar structure of hooklets and epaculates	Chironomidae	•Detailed morphological structure in Chironomidae Sp. •Identification of new structures •Understanding their functional importance	Kownacki et al. (2015)
Macaé River basin	•Distal part of maxilla bears a tuft of brush- shaped setae •Presence of short setae on the labrum	Massartela brieni Thraulodes sp.	Detailing of feeding apparatus	Baptista et al. (2006)
Wisconsim (USA)	•In male, presence of terminal lobes (2) and a post-cloacal crescent •cloacal opening is bordered with inverted U- shaped ridges hair likes structures •In female, terminal portion is cylindrical with round end and a terminal cloaca	G. difficilis	First report on intraspecific variation in body length, cuticle morphology, and gametes of this species	Bolek and Coggins (2002)
Belgium	•Pore on gnathobase surrounded by a cuticular rosette •Presence of paired gnathobase of limb bearing apical appendages (3)	Macrothrix tabrizensis	Identification of new species.	Dumont et al. (2002)
	•In male, first antenna possesses bifurcated tips of aesthetascs •Post-abdomen illustrated the glabrous end- claw.	M. agsensis
Perugia, Italy	•Pedicel exhibit sensilla chaetica delineated by an indented border •Presence of a group of sensilla basiconica •Tubular body remains surrounded by the dendritic sheath •Pedicel possess a ring of sensilla campaniformia along its distal border.	Baetis rhodani	Detailing of distribution and fine morphology of antennal sensilla of nymphal and adult mayfly	Gaino and Rebora (1998)

Figure 2.

Scanning electron micrograph depicting serrations on the inner side and presence of sensilla at the upper surface of the tarsal claws of (a) Baetis furcatus and (b) B. himalayana (Abbreviations: T-tarsus, S-spine, Cl-claw, Ug-unguitractor) (Credit: Jindal and Singh, 2020).

5. Molecular identification of macroinvertebrates

Considering the Biological Quality Element (BQE) standards, taxonomic identification displays limitations as being resource-intensive, and invasive or destructive (Hutchings, 2017; Terlizzi et al., 2003). Morphology-based identification of a species requires a large sampling size and makes the sorting of samples a bit difficult from a big pool. Importantly, getting species-level taxonomic resolution needs vast and precise taxonomic knowledge, which is both time-devouring and costly (Marshall et al., 2006). During collection, samples of macroinvertebrates can get wrecked or degrade, so the outcome of morphological identification cannot rely even upon if it has been performed by an experienced taxonomist (Haase et al., 2006). Also, morphology-based identification can be challenging due to the higher similarity in species phylogenetic makeovers (Borja et al., 2016; Costello et al., 2017).

The introduction of DNA-based identification tools can serve as a reliable, accurate, and rapid tool in identifying species or genera at any stage of life over morphological-based identification (Lobo et al., 2017; Carew et al., 2003). The utility of the molecular approach has also been observed in studies related to the identification of larvae without the availability of taxonomic keys (Brown et al., 1999; Hebert et al., 2003; Clark et al., 2001). Molecular techniques conquer those disadvantages and provide new avenues. It lowers the sample processing duration, increases the resolution, and identifies a larger number of taxa (Fayram et al., 2022).

The DNA-based technique uses precise genomic structures or “barcodes” that uniquely recognize the species (Kress et al., 2015; Elbrecht and Leese, 2017). Beermann et al. (2018b) noticed around one hundred eighty-three operational taxonomic units (OTUs) in the Chironomidae family that differentiated different species. The molecular techniques reveal species-level responses which were previously disguised by taxonomic surrogacy and provide vital information on organism response patterns w.r.t. environmental circumstances (Gleason et al., 2021). The cytochrome c oxidase I gene (CO I) region has established its role in the taxonomic identification of species by a high range of phylogenetic signals, conserved sequence, and well-known primer utility (Hebert et al., 2003; Brown et al., 1999; Clark et al., 2001; Guryev et al., 2001). Besides this, CO II and 16S rRNA are other suited regions for taxonomic identification and describing the species or genera at any stage of life (Table 2).

Table 2.

Utilization of molecular techniques and markers in the identification of macroinvertebrates.

Study site	Gene	Technique used	Species identified		References
			Family	Species
Bonn, Germany	CO I	PCR	Leptoceridae	Athripsodes albifrons	Behrens- Chapuis et al. (2021)
			Batidae	Baetis lutheri; B. niger; B. fuscatus; B. vernus Procloeon bifidum
			Caenidae	Caenis macrura
			Limnephilidae	Chaetopteryx major; C. villosa
				Potamophylax cingulatus
				Halesus digitatus
			Heptageniidae	Ecdyonurus dispar; E. insignis; E. subalpinus
			Nemouridae	Nemoura marginata Protonemura auberti
			Scirtidae	Elodes marginata
			Erpobdellidae	Erpobdella nigricollis
			Gammaridae	Gammarus fossarum
			Leptophlebiidae	Habrophlebia lauta
			Hydropsychidae	Hydropsyche incognita
			Leuctridae	Leuctra fusca
			Elimdae	Limnius opacus; L. volckmari Elmis maugeti Esolus parallelepipedus Oulimnius tuberculatus
			Polycentropodidae	Polycentropus flavomaculatus
			Asellidae	Proasellus coxalis
			Simuliidae	Simulium lineatum, S. reptans
			Pediciidae	Dicranota bimaculata;; D. gracilipes; D. pavida Pedicia littoralis
			Heptageniidae	Ecdyonurus dispar; E. insignis; E. subalpinus
			Gammaridae	Gammarus fossarum; G. pulex
			Hydraenidae	Hydraena gracilis
			Hydroptilae	Hydroptila simulans; H. sparsa
			Perlodidae	Isoperla goertzi
			Lymnaeidae	Radix balthica
			Rhyacophilidae	Rhyacophila nubile
			Sericostomatidae	Sericostoma baeticum
			Chironomidae	Tanytarsus heusdensis
South Korea	CO I	PCR	Leptophlebiidae	Choroterpes altioculus Paraleptophlebia japonica Thraulus grandis	Suh et al. (2019)
			Polmitarcyidae	Ephoron shigae
			Potamanthidae	Potamanthus formosus; Rhoenanthus coreanus
			Ephemeridae	Ephemera orientalis; E. strigata E. sachalinensis; E. separigata
			Ephemerellidae	Cincticostella levanidovae; C. orientalis Drunella aculea; D. ishiyamana D. lepnevae; D. triacantha Ephemerella atagosana; E.dentata; E. aurivillii; E. kozhovi Serratella ignita; S. setigera Teloganopsis punctisetae
			Caenidae	Caenis nishinoae
			Neoephemeridae	Potamanthellus chinensis
			Isonychiidae	Isonychia japonica; I. ussurica
			Heptageniidae	Bleptus fasciatus Cinygmula grandifolia Ecdyonurus bajkovae; E. kibunensis; E. levis; E. aesculus; E. nipponicus; E. pellucidus Heptagenia kihada; H. kyotoensis Rhithrogena japonica
			Ameletidae	Ameletus costalis; A. montanus
			Baetidae	Acentrella gnom; A. sibirica Baetiella tuberculata Baetis fuscatus; B. ursinus B. pseudothermicus; B. silvaticus; Cloeon dipterum Labiobaetis atrebatinus Nigrobaetis bacillus Alanites muticus
			Siphlonuridae	Siphlonurus chanka; S. palaearcticus
			Behningiidae	Dolania Americana
			Libellulidae	Nannophya pygmaea
				Cercion hieroglyphicum
Vellar estuary mangrove s	C O 18S rRNA	PCR	Potamididae	Telescopium Cerithidea cingulate Terebralia palustris	Thangaraj et al. (2020)
			Nassariidae	Nassarius festius
South India	COI	PCR	Baetidae	Acentrella vera Baetis sp Baetis michaelohubbardi Chopralla ceylonensis Cloeodes soldani; C. bicolor Labiobaetis soldani; L. jacobusi Nigrobaetis paramakalyani Procloeon sp. Tenuibaetis frequentus	Selvakumar et al. (2016)
			Caenidae	Caenis sp Clypeocaenis bisetosa
			Ephermerellidae	Torleya nepalica Indoganodes jobini
			Heptageniidae	Afronurus kumbakkaraiensis Epeorus petersi Thalerosphyrus flowersi
			Leptophlebiidae	Choroterpes petersi; C. nandini C. alagarensis; C. nambiyarensis Edmundsula lotica Isca purpurea Nathanella indica Notophlebia ganeshi; N. jobi Petersula courtallensis Thraulus gopalani
			Neophemeridae	Potamanthellus caenoides
			Polymitarcyidae	Languidipes corporaali
			Teloganodidae	Dudgeodes palnius Teloganodes kodai; T. sartorii; Teloganella indica
			Tricorythidae	Sparsorythus gracilis
East Asia Turkey	18S rRNA Histone H3 COI	PCR	Heptageniidae	Afronurus hyalinus; A. yoshidae Bleptus fasciatus Cinygmula grandifolia; Cinygmula sp. Ecdyonurus bajkovae; E. tigris E. tobiironis; E. viridis; E. sp. Electrogena sp. 1 Epeorus aesculus; E. curvatulus E. latifolium; E. napaeus Epeorus sp. Heptagenia kihada Proepeorus nipponicus Rhithrogena japonica; R. minazuki; R. tateyamana; R. tetrapunctigera; Rhithrogena sp.	Wakimura et al. (2016)
			Ephemerellidae	Cincticostella elongatula; C. nigra C. orientalis; C. sp Drunella cryptomeria; D. kohnoi; D. ishiyamana; D. sachalinensis; D. trispina; D. sp. Ephacerella longicaudata Ephemerella atagosana; E. notate; Ephemerella sp. Serratella setigera Teloganopsis brocha Torleya japonica Uracanthella chinoi;U. punctisetae
			Baetidae	Acentrella gnom; A. sibirica Alainites yoshinensis Baetiella japonica Baetis thermicus Labiobaetis atrebatinus orientalis L. tricolor Nigrobaetis chocoratus Nigrobaetis sp.; N. taiwanensis Tenuibaetis flexifemora
			Ephemeridae	Ephemera japonica E. orientalis; E. strigata
			Leptophlebiidae	Choroterpides nigella Paraleptophlebia chocolate P. japonica; P. spinose; P. westoni; P. sp. Thraulus grandis
			Ameletidae	Ameletus costalis; A. montanus Ameletus sp.
			Potamanthidae	Potamanthus formosus; P. idiocerus
			Siphlonuridae	Siphlonurus sanukensis; Siphlonurus sp.
			Caenidae	Caenis sp.
			Isonychiidae	Isonychia japonica
			Polymitarcyidae	Ephoron shigae
	COI	PCR	Ulmaridae	Aurelia aurita	Keskin and Atar (2013)
			Portunidae	Callinectes sapidus Carcinus aestuarii Necora puber
			Cancridae	Cancer pagurus
			Nephropidae	Homarus gammarus
			Loliginidae	Loligo vulgaris
			Majidae	Maja squinado
			Penaeidae	Marsupenaeus japonicas Melicertus kerathurus
			Mytilidae	Mytilus galloprovincialis
			Nephropidae	Nephrops norvegicus
			Octopodidae	Octopus vulgaris
			Palinuridae	Palinurus elephas
			Pectinidae	Pecten jacobaeus
			Astacidae	Pontastacus leptodactylus
			Muricidae	Rapana bezoar
			Veneridae	Ruditapes decussatus
			Scyllaridae	Scyllarides latus Scyllarus arctus
			Sepiidae	Sepia officinalis
			Squillidae	Squilla mantis
Maryland, Virginia, Pennsylvania, and North Carolina	CO I	PCR	Ephemerellidae	Ephemerella dorothea E. invaria; E. subvaria E. aurivillii; E. rotunda E. floripara; E. catawba E. excrucians; E. hispida; E. inconstans; E. infrequens	Alexander et al. (2009)
Melbourne	CO I	PCR- RFLP	Chironomidae	Ablabesmyia notibalis Cardiocladius australiensis Chironomus australis C. februarius; C. oppositus; C. cloacalis Cladopelma curtivalva Coelopynia pruinosa Cryptochironomus grisedorsum Kieffer Dicrotendipes jobetus; D. lindae Dicrotendipes sp. Kiefferulus ‘tinctus’ K. martini Microchironomus Polypedilium griseoguttatum Polypedilium nubifer Procladius paludicola Skuse Procladius spp. P. villosimanus Kieffer Rheotanytarsus trivittatus Riethia stictoptera Kieffer Tanytarsus fuscithorax	Carew et al. (2003)
			Tachinidae	Cricotopus annuliventris
New Zealand	CO I and II	PCR	Hepialidae	Wiseana copularis; W. cervinata W. fuliginea; W. jocose; W. mimica; W. signata W. umbraculata	Brown et al. (1999)
San Blas Islands		PCR- RAPDs	Briareidae	Briareum asbestinum	Cofforth and Mulawaka (1995)
			Plexauridae	Plexaura sp.; P. flexuosa P. homomalla; Pseudoplexaura porosa

As evidenced by Table 2, the molecular approach has led to the development of a large database for these macroinvertebrates. Accordingly, these have resulted in recovering almost 90% of the taxa identified morphologically and all false-absences were from taxa that represented less than 1% of organisms (GRDI-Ecobiomics, 2017).

5.1. Constrains in molecular approach

Although the molecular approach has exhibited the upper hand over morphological identification, yet not completely solved the issue (Table 3). In a metabarcoding study performed by Elbrecht et al. (2017), they reported a correct taxonomic resolution of 72.2% compared to a lower of 61.1% by morphological approach at the species level. This non-absolute identification is accredited to the non-availability of the dataset on the lesser-known families, which is a constrain posed by either the lack of the primer sequences or due to similarities in young specimens that they have left out mistakenly as replicates (Clarke et al., 2008). The cost of processing, sequencing, and expertise in DNA isolation also pose a problem, which can be easily overlooked by looking at the ease of learning, data processing, data handling (large), and less time demanding (Porter and Hajibabaei, 2018).

Table 3.

Comparative analysis of various identification techniques.

S. No.	Parameter	Morphological	Electron Microscopy	Molecular
1.	Tool used	Taxonomic keys	SEM or TEM	PCR and its types DNA barcoding
2.	Pros	•Easy to identify •Early identification •Need careful observations	•Detailing of characters •Minute characters are recorded •Obtaining spatial images of the body	•Gene amplification •Produces database •increased reproducibility and taxonomic resolution
3.	Expertise	Need not much expertise	Need expertise	Need expertise
4.	Expenses	Low	Comparatively high	Comparatively high
5.	Cons	May lead to wrong identification	•Time consuming •Needs expertise •Presence of dirt and debris	•Need complete database for accurate identification •inability to quantify taxon abundance (Barcoding) •Expensive Laboratory and sequencing costs for larger sample

6. Conclusion

Aquatic invertebrate forms like macro-invertebrates are found abundantly in water bodies as per their habitat preference associated with the physicochemical characteristics of the water. The evaluation of biodiversity and distribution of these macroinvertebrates constitute an effective tool for assessing and maintaining the physiological, chemical, and biological integrity of freshwater ecosystems. Considering the difficulty in identification or chances of misidentification, supporting molecular techniques and electron microscopy can be of great help. However, SEM and molecular approach does not present an absolute solution to the identification problem due to the high cost and need for expertise but possesses an edge over morphological studies. Molecular (DNA barcoding) studies can be improvised if backed with legacy data and with the availability of more databases along with statistical analysis.

7. Recommendations

It may include the compulsory association of molecular and ultrastructural identification tools in the identification process of macro-invertebrates. GenBank (NCBI) should be updated for lesser-known taxa and species. Alternatives methods like eDNA can be developed in order to identify life forms present in it along with water quality. Employing eDNA will prevent the loss of these aquatic forms and, single sampling will provide a comprehensive assessment of all the forms.

Declarations

Author contribution statement

All authors listed have significantly contributed to the development and the writing of this article.

Funding statement

This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

Data availability statement

Data will be made available on request.

Declaration of interest’s statement

The authors declare no conflict of interest.

Additional information

No additional information is available for this paper.