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Article

Genetic Sex Determination of Free-Ranging Short-Finned Pilot Whales from Blow Samples

by
Patricia Arranz
1,
Ruth Coya
2,
Elena Turac
2 and
Laura Miralles
2,*
1
Biodiversity, Marine Ecology and Conservation Research Group, Department of Animal Biology, Edaphology and Geology, University of La Laguna, 38200 Tenerife, Spain
2
Department of Functional Biology, Genetics Area, University of Oviedo, 33006 Oviedo, Spain
*
Author to whom correspondence should be addressed.
Conservation 2024, 4(4), 860-870; https://doi.org/10.3390/conservation4040051
Submission received: 18 October 2024 / Revised: 4 December 2024 / Accepted: 4 December 2024 / Published: 12 December 2024
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Abstract

:
Whale blow, the vapor exhaled during respiration of cetaceans, provides valuable genetic information to monitor health status and population dynamics. However, obtaining samples of sufficient quality and quantity remains a challenge, particularly for small odontocetes. Here, we developed both field and laboratory protocols optimized for the genetic analysis of blow samples of short-finned pilot whales (Globicephala macrorhynchus). Blow collection was performed from a small research vessel at a slow speed using a hand-held carbon fiber pole equipped with a sterile Petri dish. Determination of the sex was conducted using up to five PCRs of multiplexed markers from a classical methodology (SRY + ZFX/ZFY genes) and a novel protocol (SRY + FCB17) optimized for highly degraded, fragmented and/or scarce DNA. A total of 47 blow samples of free-ranging pilot whales off the Canary Islands were collected. The presence of DNA was confirmed in 98% of the blow samples, which were further processed resulting in 32 of them with positive genetic sex determination applying the novel methodology (70%), compared to only 8 (19%) with the classical method. Results confirmed the success of sampling, DNA extraction and sex determination using multiplexed markers in blow samples of odontocetes. This protocol represents an important management tool to conduct future non-invasive health assessments of small cetaceans in the wild.

1. Introduction

Cetaceans provide significant ecological contributions to the functioning of ocean ecosystems and help to mitigate climate change. Although cetaceans are umbrella species for conservation, there is still a concerning scarcity of knowledge about basic aspects of their population biology, natural history, and ecology. For most species, this impedes a correct assessment of their conservation status and their regulatory role as the largest top predators in the ocean [1].
The conservation objectives [2] included in EU directives (e.g., the Marine Strategy Framework Directive) highlight the need to improve knowledge of the biology and ecology of small odontocetes. Since these organisms spend most of their lives underwater, it is difficult to obtain reliable information about intrinsic predictor variables such as sex ratio, adult body mass, and number of births per year [3,4], which are critical aspects for assessing population health and dynamics. Animal welfare guidelines, together with technical and technological solutions available today, lead to the use of non-invasive techniques for the study of these animals, currently threatened by several anthropogenic factors (e.g., climate change, overfishing, and pollution) [2,5].
Non-invasive techniques have become a promising tool to collect data from populations to individuals without invasive procedures or causing injury. These include fecal sampling, which can be used to study diet or exposure to toxins and parasites. Environmental DNA (eDNA) from seawater is also increasingly being used to track the presence of cetaceans [6]. Both methods allow genetic analysis, but the DNA yield is low and highly fragmented [7,8]. Furthermore, it is challenging to assign the samples to a single individual, but it can be used to detect species presence [6]. In the last decade, blow sampling has been developed as a rising non-invasive technique. It is mainly used in studies about bacterial communities of whales (e.g., [9,10]), and the health status of specimens through hormonal values [11]. However, obtaining DNA of sufficient quality and quantity for genetic analyses from this type of sample is challenging, particularly for small cetaceans, for which no standardized technique has been developed yet [12,13,14,15].
Short-finned pilot whales (Globicephala macrorhynchus, hereafter: SFPWs) are odontocetes with an average size of 3.6–4.7 m in females and males, respectively [16]. SFPW females reach sexual maturity at the age of 9 and males at about 12–17 years. One single offspring is calved every five to eight years and lactated for around three years [17]. Females reach a maximum known lifespan of 60 years and remain post-reproductive in the group until death [18]. In wild SFPW populations, a balanced gender ratio (1:1) is expected, following Fisher’s principle [19], and determines the health status of a population. The male-biased sexual dimorphism in SFPWs only becomes visible from the age of sexual maturity. Until then, males and females show similar growth rates [20]. Hence, no clear visual gender assignment is possible below this age and genetics play a role in determining the sex of individuals.
SFPWs are one of the most targeted species for whale watching both globally and in the Canary Islands, where resident populations of Tenerife face several conservation challenges (i.e., marine tourism) [21]. Genetic studies on such populations are scarce, which complicates the establishment of appropriate management and conservation measures [22]. To have a non-invasive method to study these populations would help to move towards effective conservation management of the species. Considering that SFPWs log at the surface for long periods and adult specimens have a visible blow [16,23], they are a suitable model species for testing blow sampling methods. Here, a methodological approach for non-invasive genetic sample collection of free-ranging SFPWs, as well as subsequent DNA isolation and genetic sex determination, is provided, with the aim to (1) develop an optimized methodology able to estimate sex-ratio on small to medium cetaceans in the wild, (2) contribute to the long-term monitoring of the dynamics of this local SFPW population, and (3) provide a novel technique for health assessments of populations of small cetaceans in the wild.

2. Materials and Methods

2.1. Study Area and Species

Blow samples from SFPWs were collected in March and May 2024, off El Hierro (the Canary Islands, Spain), within the SAC Mar de las Calmas (ES7020057, European Union Natura 2000 Network). Data collection was performed in accordance with relevant guidelines and regulations. The research was conducted under authorization of the Spanish Ministry for the Ecological Transition and the Demographic Challenge (#AUTSPP/19/2023; #AUTSPP/33/2024). In addition, blow sampling was approved by the Ethics Committee of Animal Research and Welfare of the University of La Laguna (ref. CEIBA2021-3077).

2.2. Blow Sampling at Sea

A small research vessel (5.8 m length, Suzuki (Suzuki motor corporation, Hamamatsu, Japan) 85cv outboard engine) was used to approach SFPWs at a slow speed (<2 kn) for blow sample collection (Figure 1A). Blow sampling was performed using a hand-held carbon fiber pole (5 m length) with a sterile Petri dish (Sarstedt, Nümbrecht, Germany) of 150 × 20 mm, attached at the end with Velcro 3M Dual Lock (3M, St. Paul, MN, USA). The pole operator positioned himself at the bow of the research vessel, wearing gloves and a face mask for sterile dish manipulation (Figure 1B). The Petri dish was pointed downwards, about 50 cm above the SFPW blowhole. After the animal exhaled, the pole was retrieved and the Petri dish hand-closed to avoid seawater and airborne contamination. Petri dishes were sealed with parafilm (Laboratory film, American National Can), labeled, and stored cold until the return to port and immediately transported to the University of La Laguna (Tenerife, The Canary Islands). Once in the laboratory, samples were stored at −20 °C. Seawater samples taken from the sampling area at the time where SFPWs were present were used as environmental control samples (eDNA).

2.3. Laboratory Sample Preparation

Each Petri dish containing SFPW blow was rinsed three times with 1 mL of ethanol (absolute for analysis EMSURE®, Merck KGaA, Darmstadt, Germany) with the aid of a cell scraper Corning Incorporated (Costar©, Corning, Gilbert, AZ, USA) and then pipetted (Sarstedt; 1000 μL) into a cryovial (cryovials with external threads, silicone washer seal, self-standing, 5 mL from Simport Scientific Inc., Saint-Mathieu-de-Beloeil, QC, Canada), following that described by Atkinson et al., 2021 [15]. Care had to be taken to process one sample at a time to avoid ethanol vaporization. The lids were wrapped with parafilm (Parafilm “M”, Laboratory film, American National Can) and the vial was placed into a zip locker bag (40 × 60 mm) labeled with the sample number. Processed samples were stored at −20 °C in a dark place until shipping to the genetics’ laboratory at the University of Oviedo (Asturias, Spain).

2.4. DNA Extraction and PCR Amplification

DNA extraction was performed using the DNeasy Blood and Tissue kit, (Qiagen, Hilden, Germany), a high-efficiency extraction kit for samples including a small amount of genetic material. Prior to DNA extraction, all samples were centrifuged (4000 RCF per 10 min at 37 °C) for DNA precipitation before removing the ethanol. The resulting pellet was resuspended in 200 μL of PBS. Subsequently, DNA extraction was done following the manufacturer’s instructions with some modifications for samples with a low amount of DNA. These modifications increased the elution time (5 min), reduced the elution volume (100 μL), and performed a second elution (75 μL) through the column to increase the DNA yield. Extraction controls (only PBS, not DNA) were included.
Given the nature of blow samples and the expectation of very low DNA quantities, all procedures were done in a dedicated clean room for processing eDNA samples and inside a flow chamber to avoid any possible contamination.
To verify the presence of DNA, as well as the absence of PCR inhibitors, a generalist universal amplification was performed with the set of primers mICOIintF and jgHCO2198 [24] targeting a short fragment of 300 bp within the mitochondrial cytochrome-c-oxidase subunit I gene (miniCOI). Amplifications were performed in a conventional PCR using GoTaq® G2 Flexi DNA Polymerase Kit (PROMEGA®, Madison, WI, USA) on a VeritiPro™ Thermal Cycler (Applied Biosystems, Waltham, MA, USA) in a final volume of 20 μL containing 1× Green GoTaq® Flexi Buffer, 2.5 mM of MgCl2, 0.25 mM each dNTPs (EURx®, Gdańsk, Poland), BSA 4 mg/mL, 1 μM of each primer, 0.5 U of Go Taq G2 Flexi Polymerase, and 8 μL of sample, completed with ultrapure ddH2O. DNA was amplified following the conditions: 95 °C for 5 min, 40 cycles × [95 °C for 30 s, 50 °C for 1 min, 72 °C for 1 min] and a final extension for 7 min at 72 °C. The PCR products were then visualized on a 2% electrophoresis agarose gel with Simply Safe (EURx®, Gdańsk, Poland).

2.5. Sex Determination

2.5.1. Classical Multiplex-PCR Methodology

A multiplex-PCR was carried out by amplifying the male-specific SRY region of the Y-chromosome (~225 bp) and a fragment of the female and male zinc finger protein genes ZFX/ZFY (~445 bp) (Table 1, Figure 2), following the protocol of Jayasankar et al. [25] with modifications for samples with low amount and fragmented DNA. Modifications included increasing the sample volume to 8 μL and the addition of Bovine Serum Albumin (BSA; 4 mg/mL). Its addition reduces possible PCR inhibition and can enable successful amplification of highly fragmented DNA [26]. PCR products were visualized in ultra-pure agarose gels of 2% with Simply Safe (EURx®, Gdańsk, Poland).

2.5.2. Novel FCB17 Multiplex-PCR Methodology

A multiplex-PCR was carried out by amplifying the male-specific SRY region of the Y-chromosome (~225 bp) and FCB17 (~170 bp in SFPW—Miralles unpublished data, Table 1). FCB17 [27] is a small fragment corresponding to a cetacean short tandem repeat (STR) present in both sexes that works as a positive control in the reaction (Figure 2C). Smaller fragments are more prone to amplification in highly degraded, fragmented, or scarce DNA samples. FCB17 was first described in Delphinapterus leucas [27] but also used in Globicephala macrorhynchus [28]. Multiplexing conditions were tested in silico with the online software Multiple Primer Analyzer (https://www.thermofisher.cn/cn/zh/home/brands/thermo-scientific/molecular-biology/molecular-biology-learning-center/molecular-biology-resource-library/thermo-scientific-web-tools/multiple-primer-analyzer.html, Thermo Fisher Scientific, Waltham, MA, USA). No cross-primer dimers were detected. In silico and in vitro assessments were done prior to and after multiplexing the reaction with a set of certified male (n = 3) and female (n = 3) samples of SFPW.
Amplifications were performed in a conventional PCR using GoTaq® G2 Flexi DNA Polymerase Kit (PROMEGA®, Madison, WI, USA) on a VeritiPro™ Thermal Cycler (Applied Biosystems, Waltham, MA, USA) in a final volume of 20 μL. PCR mix included 1X Green GoTaq® Flexi Buffer, 2 mM of MgCl2, 0.25 mM of each dNTPs (EURx®, Gdańsk, Poland), BSA 4 mg/mL, 0.5 μM of both SRY primers, 0.4 μM of both FCB17 primers, 0.5 U of Go Taq G2 Flexi Polymerase, and 8 μL of the sample with ultrapure ddH2O. DNA was amplified using the following conditions: 94 °C for 1 min, 35 cycles × [94 °C for 45 s, 58 °C for 45 s, 72 °C for 1 min], and a final extension for 7 min at 72 °C. The PCR products were then visualized on a 3% electrophoresis ultra-pure agarose gel with Simply Safe (EURx®, Gdańsk, Poland).
SFPW positive controls (one male and one female) plus negative controls (DNA extraction and PCR) and eDNA samples were always added to the analyses set in both methodologies to ensure the reliability of the process. Every PCR reaction included a PCR negative control and the DNA extraction negative control. For sex determination, a PCR with all samples was five times replicated per methodology. The sample replication procedure was done for miniCOI. In total 30 negative controls were done. Sample negative control or DNA extraction negative control is a controlled sample made of distilled water instead of a blow sample. All PCR procedures were done under strict sterile conditions in a flow chamber and in a dedicated clean room for the processing of the eDNA samples.
Up to five PCR replicates were performed per sample. Each sample was assigned a reliability index (R.I.) to evaluate the confidence of the results. R.I. ranges from 1 to 4: 1—corresponding to samples in which all replicates (five) or the majority of them (four or three) had the same result; 2—two replicates showing the same result; 3—only one replicate out of five showing a result; 4—samples in which more bands than expected appeared or there was a discrepancy in the band pattern in replicates due to highly degraded or fragmented genetic material; 0—showing no results or DNA in the sample.

3. Results

In this study, a total of 47 SFPW blow samples were obtained between 7 March and 4 May 2024 off El Hierro (Table 1). All samples were processed in May 2024 and sent to the University of Oviedo for genetic analysis. Genetic analyses revealed that 46 samples (98%) had DNA when amplifying the universal miniCOI (Table 2). However, sex determination did not show positive results in all samples. Five PCR replicates were performed per sample and methodology. When using the classical multiplex-PCR methodology only eight individuals (19%) could be sexed. All samples corresponded to male individuals. Some of them only presented the SRY band, not the expected ZFX/ZFY band (Figure 2B, line M*). Whereas, when using the novel FCB17 multiplex-PCR, 32 individuals (70%) could be sexed and both sexes were represented in the samples. Only four individuals showed a multiband pattern, two of them with discrepant results. These samples were categorized as undetermined (U). R.I. scores vary from one to three in most of the samples, with three being the most frequent value. Environmental DNA controls revealed also discrepancies between methodologies (Table 1). All negative controls showed no results, as expected.

4. Discussion

In recent years, the analysis of cetacean exhaled breath or blow samples has been raising interest as a promising tool to study cetaceans in a non-invasive way (e.g., [29,30,31,32,33], and others). This study reports a novel methodology for genetic sex determination in wild SFPWs using blow samples named novel FCB17/SRY sex identification from cetacean blow.
Obtaining DNA from a cetacean blow is challenging, especially in smaller species due to lower blow volume and height [12,13]. Previous studies have reported a 21–45% success rate in the sexing of large cetaceans using blow sampling techniques [15]. However, obtaining DNA from blow samples of free-ranging delphinids promotes significant challenges [13,14] and even the absence of DNA in the collected samples, particularly due to the active behavior and small size of the species [13]. Robinson and Nuuttila [14] reported that from 37 dolphins’ blow samples, only one contained DNA. In our study with SFPWs, DNA was successfully isolated in 98% of the samples, and 70% of them were successfully sexed. This is the first report of a high success rate using blow samples for sex determination in free-ranging delphinids.
A cetacean blow contains biological fluids, mucus, lung surfactant, microorganisms, and scarce cells from the animal. Based on this complexity and the content of the sample, the presence of PCR inhibitors is highly probable, such as mucopolysaccharides, as well as DNA from the respiratory system microbiota that contain potentially many bacteria species and maybe some viruses, and scarce DNA from the target species is expected. Based on the above information, these samples have a high level of complexity with similar characteristics to eDNA samples. Therefore, in this study, blow samples were processed as eDNA, which might explain the success rate obtained. Different amplification successful rates between small universal miniCOI fragments and sex determination may be due to the possible amplification of the COI region, not only from cetacean DNA but also from microorganisms present in the respiratory biota, whereas the sex determination PCR only targets cetacean DNA. As reflected in previous studies, when considering that the extraction of microorganisms out of cetacean blow is usually successful [34,35] while sex determination of blow samples remains challenging [13,14].
Determining the sex of individuals in wild populations helps to improve knowledge of social structure, behavioral studies, and the populations’ health [36]. One of the most used genetic methods to determine the sex of cetaceans is the amplification of both the male-specific SRY gene and the ZFY/ZFX genes [37]. The detection of the SRY via PCR is the most widely used gender determination method for a wide range of mammals since this test is based on the absence of an SRY product in females due to the absence of the Y chromosome. Amplification failures of the SRY gene are likely linked to DNA degradation, a common issue in blow samples. For this reason, the ZFY/ZFX genes are also amplified along with the SRY gene in a multiplexed reaction to ensure the validity of the PCR results [25,38,39]. However, in some blow samples analyzed in this study, the amplification of the ZFY/ZFX genes failed, presumably due to DNA fragmentation, since smaller fragments such as SRY or miniCOI were positively amplified. As Konrad et al. [36] suggested, most current sexing methods are problematic with degraded DNA and samples containing degraded DNA will often fail to amplify relatively large fragments (>300 bp). To overcome this issue, a novel method with a smaller fragment (<200 bp) was proposed. Other approaches such as qPCR or restriction enzymes have been proposed in previous studies [36,38] as a solution to this problem but they require specialized equipment and are more expensive, more laborious, and time-consuming than the one we propose. Mass-sequencing might also be a possibility in large cetaceans’ blow samples but seems unfeasible in odontocetes when taking into account our results and the insights of low concentration and fragmented DNA.
When a sample presented degraded DNA and/or PCR inhibitors, multiplexed PCR reactions have been reported to fail, particularly with larger size products [40]. Smaller fragments are more prone to amplification in highly degraded, fragmented, or scarce DNA samples. For that reason, the STR FCB17 [27] of approximately 170 bp in Globicephala macrorhynchus (with only 12 bp allele range in the North Atlantic populations—Miralles unpublished data) was applied as a replacement of the ZFY/ZFX genes (~445 bp) acting as a positive control in sex determination. The STR FCB17 is a common genetic marker in odontocetes’ studies and it was used in different population genetic analyses of small cetaceans such as Delphinapterus leucas [27], Delphinus delphis [41], Stenella frontalis [41], Pontoporia blainvillei [42], and Globicephala macrorhynchus [28] but also in bigger odontocetes such as Physeter macrocephalus [43]. It was tested in blow samples of Phocoena phocoena [44] and compared with the blood samples with identical results. In this study, in silico tests confirmed the previously mentioned species and also other species such as Globicephala melas, Kogia breviceps, Tursiops truncatus, Orcinus orca, Monodon monoceros, Physeter catodon, Eubalaena glacialis, Balaenoptera musculus, and B. ricei, Eschrichtius robustus, among others. However, the list of species might be longer since in silico assessment can be only tested in complete genome data and not all cetacean species have their genome described in databases. Trials of the novel FCB17 multiplexed sex identification method in these and other species are needed to confirm its suitability. The reported tests make FCB17 a promising marker that could be extrapolated for determining the sex of blow samples from other odontocetes, particularly the challenging small species.
Cetacean blow analysis is a promising method for obtaining genetic information from populations with minimal disturbance to the animals. Here, sex determination of SFPWs from blow samples was successfully performed using the novel FCB17 multiplexed reaction. Challenges arising from the complexity of the sample were overcome by processing them as eDNA. The success of pole-sampling in SFPWs can be attributed to this species showing little reaction to the approach of slow-moving vessels, and the Petri dish can be placed close to the animal’s blowhole (<1 m), with an average of 1.8  ±  1.1 attempts per sampled whale during the first field season in 2023. This is a critical aspect of maximizing sample collection in species with a relatively small blow. While drone sampling can be challenging in small-sized species [13], it constitutes a less invasive alternative for larger cetacean species. It does not require a close approach, since the animals present a blow of several meters high, allowing the drone to fly through and their movements are more predictable at the surface [15,30]. Hence, the technique used to collect blow samples should be carefully chosen depending on the species size, behavior, and approachability of each study scenario. Further research should focus on non-invasive sampling methods to reduce disturbance and improve data collection [45] and laboratory processing. The protocols described here represent a major step towards the development of management tools to conduct non-invasive individual and population level health assessment of small free-ranging cetaceans. However, this study has some limitations such as the small sample size and restricted geographic range. Future studies with a wider geographical coverage and including more samples are suggested to confirm the results obtained. Finally, future research directions could also focus on testing the methodology in other potential species, particularly the understudied odontocetes and/or the critically endangered Balaenoptera ricei.

Author Contributions

Conceptualization, P.A. and L.M.; methodology, P.A., R.C., E.T. and L.M.; formal analysis, P.A., R.C., E.T. and L.M.; investigation, P.A. and L.M.; resources, P.A. and L.M.; data curation, P.A., R.C. and L.M.; writing—original draft preparation, P.A. and L.M.; writing—review and editing, P.A., R.C., E.T. and L.M.; visualization, P.A., R.C., E.T. and L.M.; supervision, P.A., R.C., E.T. and L.M.; project administration, P.A. and L.M.; funding acquisition, P.A. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Spanish Ministry of Science, Innovation and Universities (grant # PID2021-126940OB-I00—‘CETOCAN’ Projects for the generation of knowledge 2021) and the European Commission Horizon 2020 (grant # 0100644—‘SATURN’).

Institutional Review Board Statement

The animal study protocol was approved by the Ethics Committee of University of La Laguna (CEIBA2021-3077, 16/09/2021).

Data Availability Statement

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

Acknowledgments

We greatly appreciate the field assistance of Mónica Montoya, Elisabeth Badosa, Alicia Rodríguez, Dácil Ridolfi, Yeray Chinea, and PI of the project Natacha Aguilar de Soto. We want also to acknowledge Yaisel J. Borrell. This is a contribution from the Marine Observatory of Asturias (OMA) and the Institute of Biotechnology of Asturias (IUBA). We are also grateful to the anonymous reviewers who reviewed this manuscript.

Conflicts of Interest

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

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Figure 1. (A) Slow-speed vessel approach for blow collection of an adult male SFPW; (B) decipher SLR picture of dorsal fin for photo-ID of a blow-sampled individual. Detail of the carbon-fiber pole equipped with the sterile Petri dish used to collect whale’s blow. Photo credits: Patricia Arranz (A) and Dacil Ridolfi (B), University of La Laguna, with authorization of the Spanish Ministry for the Ecological Transition and the Demographic Challenge (#AUTSPP/19/2023; #AUTSPP/33/2024).
Figure 1. (A) Slow-speed vessel approach for blow collection of an adult male SFPW; (B) decipher SLR picture of dorsal fin for photo-ID of a blow-sampled individual. Detail of the carbon-fiber pole equipped with the sterile Petri dish used to collect whale’s blow. Photo credits: Patricia Arranz (A) and Dacil Ridolfi (B), University of La Laguna, with authorization of the Spanish Ministry for the Ecological Transition and the Demographic Challenge (#AUTSPP/19/2023; #AUTSPP/33/2024).
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Figure 2. Visualization of the different sex determination methods in agarose gel. (A) Diagram of the different fragments ordered by size and expected results (references) with a representation of the two different methods (classical and novel). (B) Results of sex determination in blow samples using the classical multiplex-PCR methodology following Jayasankar et al. [25] in 2% agarose gel. (C) Novel FCB17 multiplex-PCR proposed methodology for blow samples and highly fragmented or degraded samples in 3% agarose gel. ML, DNA Perfect Ladder 100–1000 bp (EURx®, Gdańsk, Poland); M, Male; M*, incomplete PCR result for a male; F, Female; C-PCR Negative control (no DNA).
Figure 2. Visualization of the different sex determination methods in agarose gel. (A) Diagram of the different fragments ordered by size and expected results (references) with a representation of the two different methods (classical and novel). (B) Results of sex determination in blow samples using the classical multiplex-PCR methodology following Jayasankar et al. [25] in 2% agarose gel. (C) Novel FCB17 multiplex-PCR proposed methodology for blow samples and highly fragmented or degraded samples in 3% agarose gel. ML, DNA Perfect Ladder 100–1000 bp (EURx®, Gdańsk, Poland); M, Male; M*, incomplete PCR result for a male; F, Female; C-PCR Negative control (no DNA).
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Table 1. Primer sequences for each methodology and expected amplicon size.
Table 1. Primer sequences for each methodology and expected amplicon size.
PrimerSequenceAmplicon Size
SRY forwardCCC ATG AAC GCA TTC ATT GTG TGG~225 bp
SRY reverseATT TTA GCC TTC CGA CGA GGT CGA TA
ZFX/ZFY forwardATA ATC ACA TGG AGA GCC ACA AGC T ~445 bp
ZFX/ZFY reverseGCA CTT CTT TGG TAT CTG AGA AAG T
FCB17 forwardTCA GCC TCT ATA ACG TCC TGA GC ~170 bp
FCB17 reverseATG GGG ACT GCC TAT ATT AGT CAG
Table 2. Results from the amplification of DNA and sex determination (classical multiplex-PCR and novel FCB17 multiplex-PCR proposed methodology) of individual samples, eDNA, and controls. Positive results in sex determination are in bold. Negative controls included: sample negative control (a sample made of distilled water for DNA extraction control) and PCR negative controls (the 30 negative controls performed in the PCRs).
Table 2. Results from the amplification of DNA and sex determination (classical multiplex-PCR and novel FCB17 multiplex-PCR proposed methodology) of individual samples, eDNA, and controls. Positive results in sex determination are in bold. Negative controls included: sample negative control (a sample made of distilled water for DNA extraction control) and PCR negative controls (the 30 negative controls performed in the PCRs).
Individual IDDNA
(miniCOI)		Classical Multiplex-PCRNovel FCB17 Multiplex-PCR
SexR.I.SexR.I.
24Gm01+−0F1
24Gm02+−0M1
24Gm03+−0F2
24Gm04+−0F3
24Gm05+−0−0
24Gm06+−0F2
24Gm07+−0−0
24Gm08+−0−0
24Gm09+−0F3
24Gm10+−0M2
24Gm11+−0F3
24Gm12+−0F3
24Gm13+M2M2
24Gm14+−0U4
24Gm15+−0−0
24Gm16+−0M3
24Gm17+−0U4
24Gm18+−0−0
24Gm190−0−0
24Gm20+M2M2
24Gm21+−0M3
24Gm22+−0−0
24Gm23+−0M1
24Gm24+−0U4
24Gm25+−0F3
24Gm26+M2M1
24Gm27+M2M2
24Gm28+−0M3
24Gm29+−0M3
24Gm30+−0M3
24Gm31+M2M1
24Gm32+M1M2
24Gm33+−0−0
24Gm34+−0M1
24Gm35+−0F2
24Gm36+−0M1
24Gm37+−0U4
24Gm38+−0F3
24Gm39+M2M2
24Gm40+−0−0
24Gm41+M2M2
24Gm42+−0−0
24Gm43+−0M3
24Gm44+−0M3
24Gm45+−0−0
24Gm46+−0M2
24Gm47+−0F3
eDNA C1++1+1
eDNA C2++3+1
eDNA C3+−0+1
eDNA C4++2+1
eDNA C5++1+1
Sample N.C.−−0−0
PCR N.C.−−0−0
R.I., reliability index; N.C., Negative Controls; +, positive amplification; −, negative amplification; M, male; F, female; U, undetermined; 1, samples in which the majority of replicates showed the same result; 2, samples in which two replicates showed the same result; 3, samples in which only one out of five gave a result; 4, samples in which more bands than expected appeared or showed a discrepancy in the replicates; 0, no results/DNA.
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Arranz, P.; Coya, R.; Turac, E.; Miralles, L. Genetic Sex Determination of Free-Ranging Short-Finned Pilot Whales from Blow Samples. Conservation 2024, 4, 860-870. https://doi.org/10.3390/conservation4040051

AMA Style

Arranz P, Coya R, Turac E, Miralles L. Genetic Sex Determination of Free-Ranging Short-Finned Pilot Whales from Blow Samples. Conservation. 2024; 4(4):860-870. https://doi.org/10.3390/conservation4040051

Chicago/Turabian Style

Arranz, Patricia, Ruth Coya, Elena Turac, and Laura Miralles. 2024. "Genetic Sex Determination of Free-Ranging Short-Finned Pilot Whales from Blow Samples" Conservation 4, no. 4: 860-870. https://doi.org/10.3390/conservation4040051

APA Style

Arranz, P., Coya, R., Turac, E., & Miralles, L. (2024). Genetic Sex Determination of Free-Ranging Short-Finned Pilot Whales from Blow Samples. Conservation, 4(4), 860-870. https://doi.org/10.3390/conservation4040051

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