Introduction

The super-order Xenarthra has a Neotropical distribution and is composed by two orders, Pilosa (anteaters and sloths) and Cingulata (armadillos). Xenarthra are terrestrial mammals with unique morphological attributes that are mainly related to its specialized diet and digging activities (McNab 1984; Toledo et al. 2017). The giant anteater (Myrmecophaga tridactyla), the largest representative of the living xenarthrans (Gaudin et al. 2018), is threatened by habitat loss and fragmentation, wild fire, road kill, hunting, and illegal trade (Miranda et al. 2014; Desbiez et al. 2020). The species is classified as “vulnerable” by the IUCN red list (Miranda et al. 2014) and is listed on Appendix II of the Convention on International Trade in Endangered Species of Wild Fauna and Flora-CITES. Although this vulnerable species is one of the largest terrestrial mammals in the Neotropics and plays an important role in the ecosystem, few long-term studies have focused on their ecology (Rodrigues et al. 2008; Mourão and Medri 2002; Braga 2010; Di Blanco et al. 2012; Bertassoni et al. 2020).

Giant anteater’s myrmecophagous feeding habits influence their behavior, metabolism, anatomy, and locomotor function (Naples 1999), granting them characteristics that make them challenging to anesthetize and equip with tracking devices (Rodrigues et al. 2008; West et al. 2014). This species presents a unique anatomy, with an elongated snout to accommodate its extremely long (45 cm) tongue used to reach ants and termites in their nests (Naples 1999). However, this elongated nose can make the species more susceptible to fractures during capture procedures, when inadequate methods are applied during physical restraint, especially in the wild. This species is not considered aggressive, but its long claws and strong forearms used to break termite nests have the potential to cause severe injuries to handlers during captures. Furthermore, some physiological characteristics, such as their relatively low basal metabolic rates, when compared to other placental mammals, make xenarthrans in general challenging to anesthetize (West et al. 2014).

Different capture methods and anesthetic protocols have been described for captive (Gillespie and Adams 1985; West et al. 2014; Vinci and Miranda 2012; Rojas-Moreno 2012) and wild giant anteaters (Shaw et al. 1987; Rodrigues et al. 2008; Deem and Fiorello 2002; Mourão and Medri 2002; Miranda 2004; Miranda 2008; Braga 2010; Bertassoni et al. 2020). Anesthetic remote system delivery method, such as blow-darts and dart rifles, is the most applied and described technique for giant anteaters; however, when those methods are applied associated with body weight estimation, it may increase the possibility of overdoses (Brainard et al. 2008). As a consequence, undesirable and severe effects like apnea, respiratory arrest, and hypoxia, especially in field conditions, may lead to an emergency situation, requiring assisted ventilation through chest massage, use of mask, intubation, or in the worst-case scenario, tracheotomy procedures (Brainard et al. 2008; Strom 2003). However, the xenarthran’s unique physiology grants them naturally prolonged apneas and a higher tolerance to hypoxic conditions (Deem and Fiorello 2002). In addition, their anatomy makes them more challenging species to apply assisted ventilation, such as masks, or to perform tracheal intubation, due to the limited space available to open their mouth and nasal passages (Brainard et al. 2008). Hence, monitoring of respiratory patterns or applying cardiopulmonary resuscitation (CPR) may be more complicated for xenarthran’s veterinary procedures.

Physical and chemical restraint methods should provide safety for both, animals and researchers. Hence, this study aims to evaluate and to describe physical capture and chemical immobilization protocols that enable the harnessing of free-ranging giant anteaters with GPS devices and to collect biological samples with safety and under field conditions.

Material and methods

Study site

For this study, giant anteaters were captured in four study sites, at the Pantanal (site 1) and the Cerrado (sites 2, 3 and (4) off Mato Grosso do Sul state, Brazil (Fig. 1).

Fig. 1
figure 1

Study site 1 (Pantanal wetlands), study sites 2, 3 and 4 three different highways in the Cerrado Savanna, Mato Grosso do Sul state, Brazil

Full size image

The Pantanal has a high biodiversity and is one of the largest continuous wetlands on earth (160,000 km2; Franco et al. 2013). In this pristine region, the study area was located at a private ranch in the Nhecolândia subregion of the Pantanal (Site 1, 19° 20′ S, 55°43′ W: Fig. 1). The plateau is dominated by the dry tropical Cerrado savanna, and this region is severely fragmented due to cash crops and intensive livestock farming (Reynolds et al. 2016). In the Cerrado, giant anteaters were captured in three areas (Site 2—21º 3′ S, 53º 55′ W; Site 3—21º 2′ S, 53º 54′; Site 4 –; Fig. 1).

Capture procedures

From June 2013 to July 2019, 51 wild giant anteaters were captured, and chemical immobilization was performed to collect biological samples (blood, ear biopsy, ectoparasites, feces and hair) and to fit them with GPS harnesses (TGW-4570–4 Iridium GPS) and VHF transmitters (MOD 400; Telonics, Mesa, Arizona) (Pérez et al. 2015). After approximately 1 year equipped with the harnesses, individuals were recaptured for GPS harness removal and data download.

The main method chosen for the initial capture of giant anteaters was to perform motorized active searches in open areas, using dirt roads whenever possible. Searches were performed during the winter (June-July), between 3 p.m. and 5 p.m. Anteaters present higher diurnal activity rates when exposed to colder temperatures (Di Blanco et al. 2017; Giroux et al. 2021), commonly experienced during the winter months at our study sites. Hence, performing captures during the winter facilitates the visualization and encounter of individuals and allows the capture procedures to be performed under daylight.

While driving through the study areas, two members of the team stayed in the back of the truck to improve anteater detectability. When the animal was spotted, the technicians would leave the car and approach the individual by foot. Captures were then performed using two long-handled dip-nets (handle 1.5 m; hoop 0.7 m diameter), developed and adapted by the ICAS team (Fig. 2).

Fig. 2
figure 2

Long-hand dipped nets used for physical immobilization

Full size image

Physical and chemical immobilization procedures

Two long-handled-dip-nets were used to properly restraint and keep the animals completely immobilized, hindering the animals from walking, turning, or moving their front limbs, which are equipped with long and strong claws (Figs. 3 and 4). Once the animal was properly restrained inside the two dip-nets, the veterinarian was able to safely apply an intramuscular anesthetic injection into its hind limbs (https://youtu.be/7FTNfJ64wq4).

Fig. 3
figure 3

Physical immobilization using long-hand dipped nets

Full size image
Fig. 4
figure 4

Physical immobilization using long-hand dipped nets

Full size image

The chemical immobilization protocol was based on the one described for armadillos by Kluyber et al. (2020). Here, it was applied the same combination and doses of butorphanol tartrate (Zoetis Indústria de Produtos Veterinários Ltda, São Paulo, SP, 13,064–798, Brazil; 0.1 mg/kg), detomidine hydrochloride (Agener-União, Apucarana, PR, 86,800–020, Brazil; 0.1 mg/kg), and midazolam hydrochloride (Cristália, Itapira, SP, 13,974–900, Brazil; 0.2 mg/kg), abbreviated as BDM combination.

After anesthetic induction, the first procedure before any manipulation of the individual was wrapping and completely immobilizing its front claws using 3 M™ Vetrap™ Bandaging Tape—3″ (São Paulo, SP 01315–001, Brazil). This insured the team’s safety in case the animal moved unpredictably at any time during the procedure. Then, the animal was weighted, and physical exams were performed to evaluate health conditions, detect pregnancy (palpation), general appearance, hydration status, mucous membrane color, capillary refill time, respiratory auscultation, and presence of scars or wounds. Based on this initial assessment, only adult individuals considered in good health by the team received a GPS harness. Giant anteaters that were not considered in good health were immediately released after biological sample collection and complete chemical immobilization recovery.

For anesthetic reversal procedures, all individuals received the following combination of three antagonists through deep intramuscular injection: naloxone hydrochloride (Cristália, Itapira, SP, 13,974–070, Brazil; 0.02 mg/kg) as antagonist for butorphanol tartrate, yohimbine hydrochloride (Powervet, São Paulo, SP, 05,409–010, Brazil; 0.125 mg/kg) as antagonist for detomidine hydrochloride, and flumazenil (União Química Farmacêutica Nacional, Taboão da Serra, SP, 06,785,390, Brazil; 0.01 mg/kg) for midazolam antagonism. After the end of the procedure, animals were kept in crates until complete recovery and were released at the same capture location.

Physical and chemical immobilization evaluation

Capture and chemical immobilization protocols were evaluated according to its safety, efficacy, and quality. The safety of the chemical immobilization protocol was assessed by measuring the anteaters’ vital signs every 10 min, while they were under anesthesia, until the end of all procedures. Heart rate (min; HR) was measured using a stethoscope. Respiratory rate (BPM) was measured observing and counting respiratory movements, and relative oximetry saturation (SpO2%) was measured attaching a sensor to the ear extremity or tongue—device model BN-Oxy9Vet Plus (Bionet, Tustin, CA, 92,780, USA). Finally, rectal temperature (°C; RT) was measured using a digital thermometer (Incoterm, Porto Alegre, RS, 91,751–000, Brazil).

The chemical immobilization protocol efficacy was evaluated according to the parameters proposed by Hernandez et al. (2010): induction time, total anesthetic maintenance time, and recovery time. Induction time is defined as the time from drug administration to the moment when the animal is in lateral or sternal recumbency, with lack of deep pain response, such as extremity, finger, and ear clamping. Total anesthetic maintenance time is defined as the period between the end of the induction until the first signs of voluntary motor reflexes and response to stimuli. Finally, the recovery time is defined as the period during the anesthetic procedures when the animal presents the first signs of reaction after antagonists were applied (Hernandez et al. 2010; Kluyber et al. 2020).

The chemical immobilization protocol quality was assessed for each of the three previously mentioned phases following the parameters proposed by Hernandez et al. (2010). Anesthetic induction was considered “satisfactory” when the anesthetized animal presented smooth induction and did not show signs of trembling or spasms. Anesthetic maintenance was considered “satisfactory” when the animal presented muscle relaxation upon manipulation, lack of pain and motor stimuli, and stable vital signs. Anesthetic recovery was considered “smooth” when the animal regained consciousness without excitatory movements; “disturbed” when animal regained consciousness, but presented excitatory movements (myoclonic and incoordination); and “prolonged” when complete anesthetic recovery took more than 40 min after antagonists were applied (Hernandez et al. 2010; Kluyber et al. 2020).

Statistical analysis

We estimated mean and standard deviation values to characterize each of the measured vital signs and the parameters used to assess the protocol safety and quality (Table 1). To avoid adding possible confounding effects to the analysis, first, we evaluated if the body mass of adult individuals differed between sexes using a Student’s t test. We also used Student’s t test to evaluate if average vital signs measured during the procedure (HR, BPM, SpO2%, RT) and chemical immobilization quality (induction time, total anesthetic maintenance, and recovery time) differed between adult males and females.

Table 1 Vital parameters (median ± SD) of 51 giant anteaters were anesthetized using a combination of butorphanol 0.1 mg/kg + detomidine 0.1 mg/kg + midazolam 0.1 mg/kg to allow the collection of biological samples, body measurements, and to fit GPS harness device. After about 50 ± 6.38 min of first injection of BDM combination, supplementary doses were needed (0.03 mg/kg of butorphanol, 0.03 mg/kg of detomidine, and 0.03 mg/kg of midazolam). Antagonists were applied through a combination of naloxone. 0.02 + yohimbine. 0.125 + flumazenil 0.01 mg/kg. Anesthetic stages are characterized by median and standard deviation values of: induction time, total anesthetic maintenance time, and complete recovery time after antagonists were applied
Full size table

License and authorization

All capture and handling procedures were authorized by the Brazilian Ministry of the Environment (MMA), Instituto Chico Mendes de Conservação da Biodiversidade (ICMBio), under license numberd 38,218–4 and 53,798–7.

Results

A total of 51 giant anteaters, 13 from Pantanal (site 1; 5 M, 8 F; 12 adults and 1 juvenile) and 38 from the Cerrado (sites 2, 3, and 4; 16 M, 18 F; 34 adults, 4 juveniles) were captured and chemically immobilized using the BDM protocol combination (Table 1). Average body mass of adult individuals was 32.13 ± 4.3 kg and did not differ between sexes or study areas (Table 1). The mean value of body mass, vital signs, and duration of each phase of the procedures, for each age group and sex, is presented in Table 1.

Considering both initial captures and recapture procedures to recover GPS harnesses, 130 chemical immobilizations were performed using BDM combination. The chemical immobilization protocol was considered safe and with a satisfactory quality for all of its phases. Vital signs did not differ between adult individuals of different sexes (Table 1). Cardiac rate and oxygen saturation parameters (%SpO2) oscillated during the first 20 min, but then remained stable and were not associated with clinical signs of hypoxia or hypothermia. No adverse effects like apnea, myoclonic, catalepsy, or muscle contraction were observed. Average body temperature for adults during procedures was 34.5 ± 2.52 °C. The highest average temperature of an adult during a procedure was 37.1 °C, and the lowest average was 20.2 °C (Table 1). Due to the disparate sample sizes for individuals of different age classes, we did not perform any statistical comparisons between age classes.

The BDM combination was considered effective and resulted in a quick and smooth induction for all giant anteaters captured. About 50 ± 6.38 min after induction, individuals showed signs of voluntary movements (limbs). However, a relative longer time of chemical immobilization maintenance was required for the adequate placement of GPS harnesses in individuals. Hence, supplementary doses of BDM combination were needed for procedure completion. The supplementary dose was composed of 0.03 mg/kg of butorphanol, 0.03 mg/kg of detomidine, and 0.03 mg/kg of midazolam and was applied by deep intramuscular injection in the hind limbs. Considering the supplementary doses, the mean anesthesia maintenance duration was 85.5 ± 16.8 min. Procedure duration was shorter for juveniles 71.4 ± 14.02 when compared to adults (Table 1).

At the end of each procedure, all individuals received antagonist drugs. No significant differences for recovery duration were observed between sexes (Table 1). Seven individuals received antagonist drugs intravenously (saphenous vein) and showed a faster recovery (2.55 ± 1.33 min) when compared to animals that received intramuscular injection (12 ± 5.39 min) (Table 1).

No animals died in consequence of capture, chemical immobilization procedure, or within the 24-h period after release. All giant anteaters showed signs of complete recovery up until the moment of release. No adverse reactions associated with the use of BDM combination protocol or reversal agents were observed.

Discussion

Physical immobilization

Few studies have been performed and described successful methods for capturing wild giant anteaters. Shaw et al. (1987) captured 29 giant anteaters to fit VHF harnesses by chasing the individuals and pinning them to the ground with a forked stick, and then manually injecting anesthetics. A similar method was also performed by Braga (2010) but consisted of manually holding the tail of the anteater and then applying the anesthetics. However, these methods present a great risk for field researchers if the animals escape from physical restraint before anesthetics are applied. Furthermore, anesthetic induction might be prolonged due to the animal’s stress and possibility to run.

Although giant anteaters are not considered aggressive animals, this specie’s defensive behavior using its front claws during restraint or capture procedures may cause severe human injuries (Deem and Fiorello 2002; Carregaro et al. 2009). Hence, several studies have combined the use of nets and darting, using blow pipes, for the capture of giant anteaters (e.g. Miranda 2004; Miranda 2008; Mourão et al. 2003; Bertassoni et al. 2020). However, due to giant anteater’s anatomy (with an elongated snout), the use of mesh nets can cause severe injuries to the animals (e.g., mouth trauma) and are not recommended for physical restraint. Although West et al. (2014) have recommended the darting of wild giant anteaters in the leg musculature, the authors describe intense agitation, prolonged induction, and the possibility of incomplete immobilization through this method (West et al. 2014).

Wild giant anteaters have leaner muscles when compared to captive animals, especially in their hind limbs. However, due to their long fur coat, this is not noticeable visually and might give the misleading impression of a large area to be darted. Hence, severe hind limbs injury might be caused by darting using anesthetic guns. Furthermore, screwworm infestation in the region hit by anesthetic darts has been reported (Deem and Fiorello 2002). Finally, darting using blow pipe or anesthetic guns usually requires a rough body mass estimation, and consequently increase the risk of underestimating or, especially, over-dosing the anesthesia, causing undesired effects like cardiac and respiratory depression, or apnea, and in some cases death. Hence, blow pipes and anesthetic guns are not recommended and should be avoided for giant anteaters.

The capture method developed and applied in this study, restraining animals using two long-handled dip-nets prior to anesthetic injection, was considered fast, adequate, and, most importantly, safe for the field team and the animals. None of the animals were injured during capture and handling. This method also enabled technicians to capture and completely restraint the animal before chemical immobilization. Due to the long handle of the modified dip-nets, the method used in this study avoids an unnecessarily close approach to the animal that could be risky for the team, especially during the final chasing of the individuals, before its capture (Figs. 3 and 4). Another advantage of this method is that given the initial physical immobilization of the giant anteater (i.e., with its arms and claws completely neutralized), the veterinarian can calmly choose the best location for the injection application. In contrast with the darting method, using long-handled dip-nets can guarantee the total anesthetic volume will be properly injected. As a result, applying the anesthetics, and keeping the animal restrained inside the nets, can help to promote a smoother and shorter induction, instead of chasing the animal after being darted.

Vital parameters

Due to xenarthrans’ unique physiological characteristics, such as low body temperature (30 to 36 °C) and low basal metabolic rates, previous studies suggested that anesthetic procedures should be avoided on cold days or during abrupt temperature changes to reduce the risks of hypothermia during procedures (Deem and Fiorello 2002; Miranda 2014). Nevertheless, all giant anteaters were captured for the first time during the winter (average daily temperature between 16 and 28 °C). Even though we were not able to use heating blankets under the field conditions experienced in this study, no adverse effects or significant fluctuation on body temperature during chemical immobilization were observed. Using a different drug combination, Carregaro et al. (2009) obtained lower rectal temperature (RT) parameters (32 ± 0.4 °C) for giant anteaters, when compared to this study (34.5 ± 2.52 °C).

Oximetry oscillated during the first 20 min of chemical immobilization but remained stable afterward with an average of 90.1 ± 4.38 SpO2%. The same pattern was mentioned by Carregaro et al. (2009), who also obtained low basal values (93 ± 0.5 SpO2%). However, the same authors associated these low oximetry values to the use of inhalant anesthesia, such as isoflurane (Carregaro et al. 2009), which was not used in this study. Nevertheless, Alves et al. (2020) described higher basal values for oximetry (97–99 SpO2%) obtained during one anesthetic procedure, but the anesthesia was maintained through the use of isoflurane and supplemented with 100% of oxygen 1 L/min. Despite the fact that the low basal values for SpO2 obtained in this study were similar to the values described by some authors while using different anesthetic protocols, these parameters cannot be ignored or underestimated specially during field conditions. However, studies also describe pulse oximetry (SpO2) as a popular, cheap noninvasive and portable method to detect early hypoxemia for humans and domestic animals, but its use, and accuracy, is still questionable when applied for different wild species (Mtetwa et al. 2020). Hence, the low basal values obtained from this study might also be related to the inappropriate functioning of sensor, or accuracy, or could be associated to the thick and dark tongue or ear skin, which can make oximetry reading challenging. However, no signs that could suggest hypoxia or episodes of apnea were observed during the induction, chemical immobilization maintenance, or in the end of the procedures. Although no collateral effects were observed concerning cardiorespiratory emergencies, the authors of this study highly recommend the use of supplemental oxygen especially for anesthesia of xenarthrans.

Xenarthrans are in general considered more resistant to short episodes of hypoxia (Rojas-Moreno 2014). Apnea episodes and few alterations on vital parameters have been described, and are expected, especially during giant anteater and armadillo anesthetic procedures (Rojas-Moreno 2014; Kluyber et al. 2020). The particular anatomy of giant anteaters (with a small mouth and long snout) makes them impossible to intubate, or to perform manual ventilation, and a tracheostomy would be required in case of respiratory distress (Brainard et al. 2008). To provide better accuracy and reliable data especially concerning xenarthra anesthesia, the use of portable blood gas devices is encouraged to be applied in the field, which will provide a safe and accurate monitoring and enough time to interfere or to revert an emergency event if it occurs.

Chemical immobilization protocol

To our knowledge, this is the first study to apply and describe the use of BDM combination in giant anteaters. The chosen drug combination protocol was based on the anesthetic protocols used on wild nine-banded armadillos Dasypus novemcinctus (Hernandez et al. 2010) and other four species of free-living armadillos in the Pantanal wetland, Brazil by (Kluyber et al. 2020).

Due to its cost effectiveness, dissociative agents, such as ketamine, were used to be applied alone for chemical immobilization of giant anteaters and other wild mammals; however, this is no longer recommended. Used alone, ketamine usually requires high dosages and may cause adverse effects such as respiratory depression, poor or no myorelaxation, catatonia, and a rough recovery, especially in the field (Gillespie and Adams 1985; Hernandez et al. 2010).

The second most common agent widely applied for giant anteaters and combined with ketamine are α-2 agonists, such as xylazine (Deem and Fiorello 2002; Miranda 2004; West et al. 2014). The advantages of this drug are the specific antagonism, the ability to synergize dissociative drugs, and the requirement of lower doses of ketamine. Several authors recommend this combination, stating that it provides excellent muscle relaxation, faster recovery, low cost, and high safety margins (Sinclair 2003; Hernandez et al. 2010; Rojas-Moreno 2012, 2014). However, West et al. (2014) evidenced the occurrence of severe cardiovascular abnormalities when these agents are used. Hence, medetomidine and dexmedetomidine are currently the most recommended α-2 agonist agents for giant anteaters. These drugs are chosen as an alternative to the use of xylazine, for promoting shorter analgesic effects; better muscle relaxation; quick, smooth induction and recovery; fewer adverse effects; and availability of antagonists for recovery. Furthermore, the use of medetomidine can also lower the dose of ketamine needed for anesthesia, when compared to xylazine (Rojas-Moreno 2019; West et al. 2014; Hernandez et al. 2010; Fournier-Chambrillon et al. 2000; West et al. 2014). Nevertheless, detomidine was the drug chosen for this study due to its cost effectiveness and availability.

Some α-2 agonists have been recorded to cause regurgitation and abortion (Gillespie and Adams 1985). During this study, pregnancy was detected, through palpation and baby heart auscultation by stethoscope, in four of the females anesthetized. Nevertheless, the four females were tracked by the team 2 weeks after the anesthetic procedure and were seen with newborn anteaters on their backs, suggesting that our procedure and anesthetic protocol did not cause abortions.

Similarly, to the association between α-2 agonists (xylazine, medetomidine, and detomidine) and dissociative drugs, many protocols were developed combining dissociative drugs and benzodiazepines (e.g., ketamine and midazolam or tiletamine and midazolam) for short procedures (Dahroug et al. 2009; Ferrigno et al. 2003; Rojas-Moreno 2012). This approach has been chosen especially due to the wide safety margin and excellent muscle relation promoted by this combination for xenarthras (Miranda et al. 2006; West et al. 2014; Carregaro et al. 2009). Midazolam is also recommended to avoid the risk of regurgitation, abortion (Gillespie and Adams 1985), respiratory arrest, and hypotension, effects which can be caused by xylazine (Strom 2003; West et al. 2014). Furthermore, the combination of ketamine and midazolam has been applied in free-ranging giant anteaters by Miranda (2008), and more recently by Bertassoni et al. (2020).

Opioids, such as etorphine and morphine, have rarely been used in giant anteaters and have generally been used as pre-anesthetics, combined or not with benzodiazepines such as diazepam. However, they are considered impractical for field situations (Deem and Fiorello 2002; West et al. 2014). Butorphanol was chosen for this study because it has been commonly used and recommended for other wild animals’ anesthesia, e.g., tapirs and armadillos (Hernandez et al. 2010; Medici et al. 2014; Kluyber et al. 2020). Usually combined with other anesthetic drugs, butorphanol can also be applied for pain control after surgical procedures and even to reduce the volume or dosage required for other drugs (Hernandez et al. 2010).

At the end of the procedures and after antagonists were applied, animals were kept in wooden crates, until they demonstrate signs of complete anesthesia recovery. In contrast with what was described by West (2014), keeping giant anteaters inside wooden boxes provided them a smooth recovery and avoided animals to be injured due to recovery excitement especially after antagonist agents were applied.

Safety and advantages

To our knowledge, this study counts on the largest number of wild giant anteaters ever captured and anesthetized. An unprecedented total of 130 chemical immobilizations (considering first capture and recapture to remove the collars) were performed using the proposed anesthetic protocol. Furthermore, this study is part of a long-term xenarthran conservation, ecology, and health assessment initiatives performed by the Institute for the Conservation of Wild Animals since 2011. Within this initiative, 300 chemical immobilizations and anesthesia procedures have been safely performed using the BDM combination protocol, in several free-living xenarthran species including giant armadillos (Priodontes maximus), nine-banded armadillos (D. novemcinctus), six-banded armadillos (Euphractus sexcinctus), southern naked-tailed armadillos (Cabassous unicinctus) (Kluyber et al. 2020), southern tamanduas (Tamandua tetradactyla), and giant anteaters (D. K. unpublished data).

One of the main advantages of this protocol is that it can be completely reversed at any time during the procedure, promoting smooth, rapid recoveries. Hence, the use of antagonist drugs provided a key contingency plan in case of emergency or adverse collateral effects. Nevertheless, no adverse effects or emergency situations were faced during the procedures due to the use of specific anesthetic drugs or its reversal agents (e.g., excessive salivation, vomiting, regurgitation, or bloating). Specialized equipment was not required during procedures. No animals died upon capture or before, during, or within 24 h after the procedure, and all the animals were completely recovered at the time of release.

This protocol provided rapid, smooth inductions and a wide safety margin. The large therapeutic window and low cost of this combination per procedure/individual makes this method an attractive choice in environments where monitoring and resources are scarce or present difficult logistic, especially for emergency procedures or when compared to zoos or hospital facilities. Hence, based on the results of this study, the proposed immobilization protocol was considered efficient, safe, and highly feasible, especially under field conditions. This study reinforces the safety of performing the chemical immobilization of xenarthrans through the BDM drug combination protocol and recommends its use for giant anteaters and other xenarthran species.