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Buccal jet streaming and dead space determination in the South American lungfish, Lepidosiren paradoxa
Comparative Biochemistry and Physiology, Part A 235 (2019) 159–165
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This article is part of a Special issue on Physiology from the Neotropics
Buccal jet streaming and dead space determination in the South American lungfish, Lepidosiren paradoxa☆ Walter J. Mintoa, Humberto Giustia, Mogens L. Glassa,1, Wilfried Kleinb,c, Glauber S.F. da Silvac,d,
T ⁎
a
Faculty of Medicine of Ribeirão Preto, University of São Paulo, Ribeirao Preto, SP, Brazil School of Philosophy, Sciences and Literature of Ribeirão Preto, University of São Paulo, Ribeirão Preto, SP, Brazil c National Institute of Science and Technology on Comparative Physiology, Rio Claro, SP, Brazil d Institute of Biological Science, Department of Physiology and Biophysics, Federal University of Minas Gerais, Belo Horizonte, MG, Brazil b
A R T I C LE I N FO
A B S T R A C T
Keywords: Buccal pump Air-breathing Lung gas End-tidal CO2 Effective ventilation Bohr equation Lungfish
The “jet stream” model predicts an expired flow within the dorsal part of the buccal cavity with small air mixing during buccal pump ventilation, and has been suggested for some anuran amphibians but no other species of air breathing animal using a buccal force pump has been investigated. The presence of a two-stroke buccal pump in lungfish, i.e. expiration followed by inspiration, was described previously, but no quantitative data are available for the dead-space of their respiratory system and neither a detailed description of airflow throughout a breathing cycle. The present study aimed to assess the degree of mixing of fresh air and expired gas during the breathing cycle of Lepidosiren paradoxa and to verify the possible presence of a jet stream during expiration in this species. To do so, simultaneous measurements of buccal pressure and ventilatory airflows were carried out. Buccal and lung gases (PCO2 and PO2) were also measured. The effective ventilation was calculated and the dead space estimated using Bohr equations. The results confirmed that the two-stroke buccal pump is present in lungfish, as it is in anuran amphibians. The present approaches were coherent with a small dead space, with a very small buccal-lung PCO2 difference. In the South American lungfish the dead space (VD) as a percentage of tidal volume (VT) (VD / VT) ranged from 4.1 to 12.5%. Our data support the presence of a jet stream and indicate a small degree of air mixing in the buccal cavity. Comparisons with the literature indicate that these data are similar to previous data reported for the toad Rhinella schneideri.
1. Introduction The six species of extant Dipnoi (Neoceratodus forsteri; Lepidosiren paradoxa; Protopterus aethiopicus, P. amphibius, P. annectens, P. dolloi) possess lungs as their principal air-breathing organs (Graham, 1997). The morphological characteristics of dipnoan lungs have been studied for centuries (Owen, 1841; Quekett, 1844; Hyrtl, 1843; Günther, 1871; Parker, 1892; Poll, 1962; Grigg, 1965; Klika and Lelek, 1967; Hughes and Weibel, 1976; Maina and Maloiy, 1985; de Moraes et al., 2005). Several aspects of lung ventilation, gas exchange, blood gas chemistry, or chemoreception have also been of interest for comparative physiologists, especially over the last 50 years (e.g. Johansen and Lenfant, 1967; Johansen and Lenfant, 1968; Lenfant et al., 1970; McMahon, 1970; Fritsche et al., 1993; Amin-Naves et al., 2004; DeLaney et al., 1983; Sanchez and Glass, 2001; Glass, 2010; Sanchez and Glass, 2001;
Sanchez et al., 2001b; Bassi et al., 2005; Bassi et al., 2010; Sanchez et al., 2005; Amin-Naves et al., 2007a, 2007b; da Silva et al., 2008, 2017; Zena et al., 2017). Such attention given to morphology and physiology of lungfish can be explained by their phylogenetic position close to the origin of the Sarcopterygii (Brinkmann et al., 2004; Yokobori et al., 1994; Zardoya et al., 1998) and their ability to exchange gases bimodally with water (across gills and/or integument) and air (using lungs), representing the remaining living species of a lineage of vertebrates that evolved air breathing in the mid-Devonian (~380 Ma; Clement and Long, 2010). One aspect of lungfish respiration, however, has not received much attention up to now: the breathing mechanism (Bishop and Foxon, 1968; McMahon, 1969; Brainerd, 1994). While studies of the buccal pump mechanism have been carried out on L. paradoxa (Bishop and Foxon, 1968; Brainerd, 1994) and P. aethiopicus (McMahon, 1969;
☆
This article is part of a special issue entitled: Physiology from the Neotropics, edited by: Dr. Kenia Bicego, Dr. Luciane Gargaglioni and Dr. Mike Hedrick Corresponding author at: Departamento de Fisiologia e Biofísica, Instituto de Ciências Biológicas; Universidade Federal de Minas Gerais, Av. Presidente Antonio Carlos, 6627. ICB/UFMG, Depto Fisiologia e Biofísica. Sala 158, Bloco B4-158, Pampulha, 31270-901 Belo Horizonte, MG, Brazil. E-mail address: [email protected] (G.S.F. da Silva). 1 In memorian, Deceased October 4th, 2018. ⁎
https://doi.org/10.1016/j.cbpa.2019.05.026 Received 3 March 2019; Received in revised form 8 May 2019; Accepted 20 May 2019 Available online 10 June 2019 1095-6433/ © 2019 Elsevier Inc. All rights reserved.
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2. Material and methods
Brainerd et al., 1993), only superficial descriptions of breathing behavior are available for N. forsteri (Dean, 1906; Grigg, 1965). It seems clear, however, that all lungfish use a two-stroke buccal pump, composed of one expiration and at least one inspiration, to renew the air within the lungs (Brainerd, 1994). The events during one breathing cycle can be described as follows (Brainerd and Ferry-Graham, 2006): 1) Air breathing starts with the animal putting its head out of the water and the mouth being opened; 2) The buccal cavity expands to draw in oxygen-rich air; 3) As the buccal cavity expands, the glottis opens and the lungs empty through elastic recoil of body wall and lungs; 4) Once the buccal cavity has been fully expanded, the mouth closes and the air trapped within the buccal cavity is pushed into the lungs. Since fresh air and gas expired from the lungs enter the buccal cavity during buccal expansion, some degree of mixing of oxygen-rich and oxygen-poor gas occurs during the first phase of the two-stroke buccal pump, reducing the amount of oxygen entering the lungs. McMahon (1969) estimates the gas mixing to be 20–40% of expired air being returned to the lungs in P. aethiopicus. A two-stroke buccal pump ventilation has also been identified in anuran amphibians. De Jongh and Gans (1969) and Gans et al. (1969) performed classical experiments using the bullfrog (Lithobates catesbeianus) to describe the complex functioning of the buccal pump. De Jongh and Gans (1969) identified three different types of cyclical phenomena associated to buccal movements. 1) Oscillations of the buccal floor to ventilate the buccal cavity through the open nares. 2) Ventilatory cycles to renew the air within the lungs through coordinated opening and closing of nares and glottis. 3) Inspiratory movements with pulmonary ventilation interspaced by non-ventilatory periods. One fundamental proposal of de Jongh and Gans (1969) was the presence of an expiratory airflow, known as jet stream, postulated to result in minimal mixing of oxygen rich air in the buccal cavity with the oxygen poor air leaving the lungs during expiration, and passing rapidly along the dorsal part of the buccal cavity. For Lithobates pipiens a similar ventilatory pattern as in the bullfrog has been described, but no evidence for the presence of an expiratory jet stream has been found (Vitalis and Shelton, 1990), since the air being pumped into the lungs during inspiration represents a mixture of air, containing 30–50% of air from the previous expiration (West and Jones, 1975). Brett and Shelton (1979), studying Xenopus laevis, found in this species a ventilatory cycle starting with an expiration that is followed by an inspiration, thereby suggesting no mixing of fresh and used air within the buccal cavity. Fernandes et al. (2005), investigating the ventilatory pattern and jet stream hypothesis in the toad Rhinella schneideri, found a buccal cavity dead-space representing 30–40% of tidal volume, suggesting some degree of inspiratory and expiratory flow separation, with limited mixing within the buccal cavity. Regarding the ventilatory cycle of L. paradoxa, it has to be assumed that this species possibly expires almost all of the air contained within the lungs at the beginning of a ventilatory cycle. This was demonstrated by Bishop and Foxon (1968) using X-ray imaging of a breathing cycle, where the lungs were no longer visible during expiration due to a nearcomplete loss of air, also has been suggested by da Silva et al. (2017), based on measurements of expiratory tidal volume. Since pulmonary dead space seems to be very small, a significant amount of gas mixing may only be possible during the expansion phase of the two-stroke buccal pump, when air is being expired from the lungs and fresh air is being drawn into the buccal cavity. To determine the degree of mixing of fresh air and expired gas during the breathing cycle of L. paradoxa and to verify the possible presence of a jet stream during expiration in this species, we performed measurements of buccal pressure, buccal and pulmonary PCO2 and PO2, pulmonary ventilation and calculated dead space and effective ventilation by using the Bohr dead space equation (see Eq. 3).
2.1. Animals The lungfish specimens of Lepidosiren paradoxa were collected in the region of Cuiabá – MT (Brazil) and transported to the University of São Paulo, campus Ribeirão Preto (FMRP/USP) (authorized by IBAMA # 02027. 002172/2005–68). Upon arrival, the animals were placed in tanks of 1000 L with aerated and temperature-controlled water (25 °C) and were fed three times a week with chicken liver. Before the experiments, each animal remained 24 h without access to food. The experimental procedures were approved by the local ethics and animal care committee (Proc. # 0762005). Ten animals (N = 10) of both sexes were used in the present study. In order to avoid running experiments during the aestivation period (dry season – from May to September) all procedures took place during animal's active season, from October (2011) to April (2012).
2.2. Surgical procedures Animals were anesthetized by immersion in a solution of benzocaine to 1:1 g/L as extensively described in our previous studies (Amin-Naves et al., 2007a, 2007b; da Silva et al., 2011; da Silva et al., 2008). Under anesthesia, lungs and buccal cavity were cannulated in different groups of animals. For cannulation of the lung, anesthetized animals were transferred to the surgical apparatus, in which anesthesia was maintained by switching to a diluted (1:4) solution of benzocaine with water through a catheter onto the gills. An incision (~2 cm) was made into the body wall in the dorsal caudal region to expose the posterior part of the lungs. Care was taken not to damage any closely positioned vessels. Then, a cannula (PE 50 connected to a PE 90) was inserted into the lung through a small incision and secured externally around the lung. The catheter was exteriorized and the skin sutured. The buccal cavity was cannulated using a polyethylene cannula (PE 50) connected to a PE 90, which was inserted into the buccal cavity through a micro incision in the lateral buccal floor. This cannula was used to collect air samples (for PO2 and PCO2 measurements) and to record pressures. After surgical procedures, animals were allowed to recover for at least one day before starting the experimental protocols.
2.3. Measurements of pulmonary ventilation The experimental apparatus used for pulmonary ventilation recordings was the same as described in our previous studies (da Silva et al., 2011, 2017). Briefly, animals were placed into a closed plastic box that contained perforated sidewalls for water circulation. An inverted funnel was attached to the lid of the plastic box, representing the only place for the animal to surface to breathe air. A homemade pneumotachograph (formed by a set of parallel tubes) was attached to the funnel to record ventilatory airflow. This setup was completely immersed into a much larger Styrofoam box and a submerged pump, which was placed outside of the plastic box, was used to maintain water circulation within the experimental apparatus. Animals were placed into this apparatus for a period of 12 h for acclimatization before starting the experiment. Pulmonary ventilation was measured using the pneumotachographic method described by Glass et al. (1983) for aquatic animals. A differential pressure transducer (141 ML spirometer; ADInstruments, Sydney, Australia) was connected to the pneumotach and recorded breathing-related pressure oscillations. The system was calibrated by injecting and withdrawing a known volume of air (10 mL). This method was used for determining air-breathing frequency (fR) and tidal volume (VT, mL kg−1). The calculation of total lung ventilation (VĖ TOT, mL BTPS Kg−1 h−1) was obtained by multiplying fR by VT. 160
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as a percentage of tidal volume (VT) (Fernandes et al., 2005):
2.4. Experimental protocols
VD/ VT = (PL CO2 –PB CO2)/ PL CO2
All the experiments were carried out during the active season, daytime at normoxic (20.95% O2) and normocarbic (0.03% CO2) conditions (at both gas and water phase) and at controlled temperature of 25 °C.
(3)
where: PBCO2 represents the CO2 partial pressure in the buccal cavity during a breathing episode. This calculation is usually based on intrapulmonary PCO2 (PLCO2) measurements compared to end-tidal PCO2 (PETCO2) during expiration of the breathing episode (Fernandes et al., 2005). However, it is not possible to place a mask onto the lungfish in order to measure end-tidal gas compositions, thus we used PBCO2 as substitute for PETCO2. Similar to alveolar ventilation (V̇ A), the effective pulmonary ventilation (V̇ EF) was obtained as (Wang et al., 1998; Fernandes et al., 2005):
2.4.1. Measurements of buccal pressure To measure variations in buccal pressure associated to the ventilatory cycle, an animal was placed into the experimental apparatus and had the buccal catheter connected to a pressure transducer (MLT0380; ADInstruments, Sydney, Australia). The transducer signals passed through a single-channel Bridge Amp (FE221, ADInstruments) and were recorded using a data acquisition system (PowerLab 4/30; AdInstruments) and analyzed using LabChart 7 software (AdInstruments).
V̇ Eff = (V̇ CO2•RT )/P LCO2
(4)
where: V̇ Eff gives the effective ventilation of the lung, V̇ CO2 the CO2 elimination through the lung, PLCO2 the intrapulmonary (lung) PCO2, R the gas constant, and T the absolute temperature (K). V̇ CO2 was not measured in the present study and hence the value used to calculate V̇ Eff was obtained from our previous article (da Silva et al., 2017). Knowing V̇ Eff and V̇ ETot, dead-space was also calculated as:
2.4.2. Measurements of partial pressure of CO2 (PCO2) and O2 (PO2) in lung and buccal compartments The purpose of this protocol was to obtain the partial pressure of CO2 and O2 within the buccal cavity and lungs of unanaesthetized L. paradoxa. Animals within the experimental apparatus had the pulmonary cannula connected to a 5 mL syringe. Firstly, to avoid any collapsed areas within the lungs, a volume of 5 mL of room air was injected. Then, the same volume of air was withdrawn from the lung after four different time-points (1, 10, 20 and 40 min). The lung air samples were injected into a gas analyzer (ML206 ADInstruments, Sydney, Australia) and the gases were recorded using the same data acquisition system as for the pressure measurements. Importantly, the syringe was sealed using a three-way stopcock to avoid any leaking and minimize any possible mixture with room air. Air samples from previously mentioned time points were averaged in each animal and considered for the further calculations, where values of lung PO2 and PCO2 (PLO2 and PLCO2, respectively) were needed. The gas composition within the buccal cavity was measured through the buccal catheter being connected to the gas analyzer and data acquisition system (ADInstruments). A small sampling flow rate was obtained by the analyzer built-in pump. The oscillations in O2 and CO2 were monitored to determine gas partial pressures in the buccal cavity (PBO2 and PBCO2) during spontaneous air breathing (ventilatory movements). To accomplish these measurements, the animal was kept not totally submerged in water, but rather, in an aquarium with a thin layer of water, enough to keep it humidified. This procedure allowed us to perform buccal gas sampling without any water from the buccal cavity being sucked to the gas analyzer. Gas recordings were obtained as percentage (%) or fraction and were subsequently converted to mmHg considering a barometric pressure of 710 mmHg and the partial pressure of water vapor at 25 °C.
̇ –VEff ̇ )/ fR VD = (VETot
(5)
Knowing tidal volume (VT), the relationship VD / VT was obtained. Lung and buccal PO2 and PCO2 were compared using Student's ttest. All data are reported as mean and standard error (mean ± SE). Significance level was P < .05. The N-value was 5 for each data set. 4. Results Fig. 1 depicts a representative buccal pressure profile along with airflow measurements during a breathing event in the lungfish. A
3. Equations used and statistics The lung O2 extraction coefficient (EO2) (which expresses the percentage of O2 absorbed by the pulmonary capillaries) and the respiratory exchange ratio (RER) were calculated following (Dejours (1981):
E O2 (%) = [PI O2 − PL O2)/ PI O2]•100
(1)
RER = PL CO2 /(PI O2 − PL O2)
(2)
Fig. 1. Measurements of buccal pressure (A) and ventilatory air flow (B) in L. paradoxa at 25 °C. The letters ‘a’ to ‘k’ represent events associated to changes in buccal pressure and airflow. ‘a’ represents the start of a ventilatory cycle followed by a decrease in buccal pressure to sub-atmospheric values; ‘b’ marks initiation of expiratory flow; ‘c’ represents the increase in buccal pressure associated with air being pushed from the buccal cavity into the lungs; ‘d’, ‘f’, ‘h’ give peak buccal pressures and mark initiation of inspiratory airflow; ‘e’, ‘g’, ‘i’ represent inspiratory airflow and the initial increase of buccal pressure associated with air being pushed into the lungs; ‘j’ marks the last peak buccal pressure and flow measurements returning to zero flow; ‘k’ marks buccal pressure oscillations associated to the elimination of remaining air from the buccal cavity.
where: PIO2 gives the oxygen partial pressure of inspired air (in mmHg), PLO2 the oxygen partial pressure in the lungs (in mmHg), and PLCO2 the lung carbon dioxide partial pressure (mmHg). Eq. 2 is a simple rearrangement of the alveolar gas equation, where alveolar PO2 and PCO2 (PAO2 and PACO2) were replaced by the measured values of PLO2 and PLCO2. The Bohr dead-space equation was used to estimate dead space (VD) 161
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Table 1 Respiratory variables measured and calculated in the lungfish L. paradoxa in the present study. Respiratory variables
Mean ± SE
V̇ E (mL BTPS h−1 kg−1) Effective V̇ Eff (mL BTPS h−1 kg −1) VT (mL kg−1) fR (breaths h−1) Lung PCO2 (mmHg) Buccal PCO2 (mmHg) Lung PO2 (mmHg) Buccal PO2 (mmHg) RER Extraction Coefficient (%) EO2 (%) = [PIO2-PLO2)/PIO2] •100
89.85 ± 9.70 82.77 ± 5.80 27.08 ± 2.80 2.0 ± 0.2 20.63 ± 1.60 19.90 ± 2.40 93.80 ± 6.80 87.40 ± 11.10 0.42 ± 0.04 35.3 ± 4.7
PCO2 and PO2 values were statistically not different in the lung and buccal compartments (PCO2; buccal vs lung: p = .8 and PO2; buccal vs lung: p = .64). Table 2 Calculation of dead-space in percentage of tidal volume (VD/VT) and comparison between the South American lungfish L. paradoxa and the Toad Rhinella schneideri.
Fig. 2. Values of gases partial pressures in the lung (circles) and buccal (squares) compartments. PCO2 values are depicted in panel A and PO2 in the panel B. Each point represents the value of an individual animal, while horizontal lines (middle and top-bottom) represent mean and standard error. There is no statistical difference in the PCO2 and PO2 between lung and buccal compartments (N = 5).
1) VD = (VTOT – VEff) / fR
Lungfish
Toad
Effective ventilation (mL BTPS •min−1 kg −1) Total ventilation (mL BTPS •min−1 kg −1) Breathing frequency (cycles •min−1) Tidal volume (VT) (mL •kg−1) Dead space volume (VD) (mL •kg−1) (VD/VT) 100%
1.37 1.48 0.03 27.0 3.54 12.5
7.00 12.10 5.80 2.4 0.90 37.0
2) VD/VT = (PLCO2 – PBCO2) / PLCO2 PLCO2 (mmHg) PBCO2 (mmHg) (VD/VT) 100%
20.6 19.9 4.12
10.9 7.6 30.2
3) Substitution of PLCO2 by PaCO2 PaCO2 (mmHg)⁎ (VD/VT) 100%
21.8 8.7
12.3 49.9
⁎ PaCO2 value has been calculated as average from several studies on the lungfish L. paradoxa at 25 °C in air and water normoxia (for details see review article submitted). For purposes of comparison, the numbers presented for both, lungfish (present study) and toad (Fernandes et al., 2005) are mean values.
typical ventilatory cycle in L. paradoxa was characterized by a slight increase in buccal pressure (‘a’ in Fig. 1) followed by a decrease in buccal pressure to sub-atmospheric values. Once negative pressure stopped decreasing, expiratory flow initiated (‘b’) and continued for about 4 s. With expiratory flow still occurring, buccal pressure increased sharply to positive values (‘c’). After reaching a peak buccal pressure of about 6 mmHg (‘d’), pressure in the buccal cavity fell to negative values again, coinciding with an inspiratory flow. Buccal pressure rose again sharply for another three times to positive values (9–12 mmHg), always accompanied by a decreasing respiratory flow (‘e’, ‘g’, ‘i’). Following each of these increasing buccal pressures were three sharply decreasing buccal pressures (‘f’, ‘h’, ‘j’), that were coincident with inspiratory air flows, with the exception of the last decrease in buccal pressure (‘j’). At this moment (‘j), ventilatory flow measurements returned to zero flow and buccal pressure oscillated between positive and negative values before returning to zero. All compressions and expansions of the buccal cavity (from ‘c’ to ‘k’) took 0.8 to 1.0 s each. Partial pressures of O2 and CO2 from the lungs and the buccal cavity are given in Fig. 2 and Table 1 and the mean values of V̇ E, VT and fR, along with other measured and calculated respiratory variables are presented in the Table 1 (all under normoxic conditions and at 25 °C). The PaCO2 and V̇ CO2 values were taken from previous studies (review article; da Silva et al., 2017) and used to calculate effective ventilation (eq. 4), which corresponded to a large fraction of the total ventilation. This result agrees with the relatively small dead-space (VD) found. Although lung PO2 and PCO2 tended to be greater than values measured in the buccal cavity, no statistical differences were found between lung and buccal gas composition (Fig. 2). The O2 extraction
coefficient, calculated using the inspired-lung PO2 difference (eq. 1), was found to be 35.3 ± 4.7%. Table 2 demonstrates the calculations of dead-space in percentage of tidal volume (VD / VT) and comparisons between the South American lungfish L. paradoxa and the toad Rhinella schneideri as described by Fernandes and et al. (2005). Using the total ventilation, effective ventilation and breathing frequency, the VD / VT ratio was estimated to be 12.5% (eq. 5). In absolute values, the deadspace was calculated and estimated to be 3.5 mL kg−1. Based on the small difference in lung and buccal PCO2, the dead-space calculated using the Bohr equation (eq. 3) corresponded to 4.1 ± 6% of the tidal volume. Substitution of lung PCO2 by PaCO2 gave a dead-space of 8.7% of tidal volume. Thus, in the South American lungfish the VD / VT ranged from 4.1 to 12.5%, which were smaller than those values reported for R. schneideri. 5. Discussion Pulmonary airflow measurements in the current study confirm the presence of a two-stroke breathing mechanism in L. paradoxa, composed of a single expiration followed by 2–4 buccal inspirations. Our recordings of buccal cavity pressure, however, were different from previous recordings, since a negative buccal pressure was found at the beginning of each ventilatory cycle, always coinciding with expiratory flow. While McMahon (1969) and Brainerd et al. (1993) did record in P. aethiopicus small negative pressures in the buccal cavity at the 162
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Table 3 Mean values reported in the literature for pulmonary ventilation (V̇ E, ml BTPS kg−1 h−1), tidal volume (VT, ml BTPS kg−1), and breathing frequency (fR, breaths h−1) in L. paradoxa at 25 °C.
Fig. 3. Postulated airflow pattern during the first expansion of the buccal cavity in Lepidosiren paradoxa in frontal view of the open mouth (A) and in lateral view (B). Blue traces in A show the postulated jet stream of air along the dorsal part of the buccal cavity, being guided by dorsal folds towards the posterior internal nares and exiting through the anterior internal nares. Blue arrows in B show expiratory airflow leaving through the anterior internal nares, while the red arrow represents inspiratory flow. Panel A modified from Hyrtl (1843) and panel B modified from Goeldi (1898).
Reference
V̇ E
VT
fR
Amin-Naves et al. (2004) Amin-Naves et al. (2007a) Amin-Naves et al. (2007b) Bassi et al. (2010) da Silva et al. (2011) da Silva et al. (2017) Harder (2001) Sanchez et al. (2001a) Sanchez et al. (2001c) Sanchez et al. (2005) Present study
162.0 22.1 39.0 63.7 64.9 73.4 42.0 88.2 103.0 43.0 89.9
29.0 15.0 20.0 20.5 29.8 27.3 27.8 24.0 26.0 28.0 27.1
5.4 1.0 1.3 3.2 2.1 2.6 1.4 5.5 4.0 1.7 2.0
Similarly, the data obtained regarding lung PCO2 and PO2 are in the same range as in previous studies (Johansen and Lenfant, 1968; Bassi et al., 2005; da Silva et al., 2011). Fernandes et al. (2005), using identical methods to calculate ventilatory dead space in Rhinella schneideri, found dead space to make up between 30 and 50% of VT. No other estimates for ventilatory dead space are available for amphibians, nor any air-breathing fish. A comparison with amniotes seems not valid, since their respiratory system is composed of lungs being connected through a trachea to the mouth cavity, a morphological feature not seen in any air-breathing fish and amphibian. In lungfish, the lungs are connected through a short duct to the esophagus, and this connecting duct is not maintained open with cartilaginous rings as in an amniote trachea and remains closed between pulmonary breathing cycles. Therefore, only the volume of the buccal cavity and any residual air in the lungs contribute to respiratory dead space in a lungfish. Our calculations suggest dead space volume in L. paradoxa being between 4.1 and 12.5% of VT, values significantly lower than in R. schneideri. These low values may be due to the relatively small buccal cavity with a relatively constant volume when compared to the toad. Da Silva et al. (2017) estimated maximum buccal breath volume to be around 15 mL kg−1 in L. paradoxa, a value much smaller than the 20–50 mL kg−1 reported for Lithobates catesbeianus (Gans et al., 1969) or the estimate of 25 mL kg−1 for R. schneideri (Fernandes et al., 2005). Assuming a mean dead space volume of 10% for L. paradoxa and a buccal cavity volume of 15 mL kg−1, only a volume of 1.5 mL kg−1 would constitute the dead space in this species. This small dead space volume would only be returned towards the lungs during the first buccal inspiration, whereas the following buccal inspirations would each pump 100% fresh air into the lungs. Assuming that lungfish exchange about 90% of pulmonary air during each ventilatory cycle (Bishop and Foxon, 1968; Brainerd, 1994; Harder, 2001; da Silva et al., 2017), a one kilogram animal would possess a lung volume of 37 mL, from which 33.3 mL would be expired and accordingly 3.7 mL would remain with the lungs. From the exhaled 33.3 mL, 1.5 mL would be mixed with inspired air and returned to the lungs. Consequently, immediately after a ventilatory cycle the lungs would contain 5.2 mL of used air and 31.8 mL of fresh air. Assuming a PIO2 of 150 mmHg and a PLO2 of 30 mmHg after a 10 min breath-hold, PLO2 immediately after inspiration and full mixture of pulmonary air, would be about 133 mmHg. And indeed, Johansen and Lenfant (1967) report PLO2 to be immediately after a ventilatory cycle at up to 140 mmHg in L. paradoxa and McMahon (1970) reports PLO2 of above 110 mmHg in P. aethiopicus, suggesting a nearly complete renewal of air during the two-stroke buccal pump mechanism. Moreover, considering that the Bohr VD equation deals with dilution of gas volumes, our data suggest that the degree of gas mixing within the buccal cavity was limited. If the following assumptions are considered: (i) buccal cavity volume ~ 15 mL kg−1, (ii) the gas expelled from the lungs mixes with the buccal gas, (iii) During non-ventilatory periods and immediately before
beginning of or before an expiration (corresponding to an opening of the mouth at the beginning of the ventilatory cycle; Brainerd et al., 1993), as well as immediately after a first buccal inspiration, they were not seen during expiratory airflow. We recorded a constant negative pressure being maintained within the buccal cavity (Fig. 1), and the accompanying ventilatory recordings are similar to the constant outflow of air seen in a previous study (da Silva et al., 2017). The presence of a constant negative pressure simultaneously to expiratory airflow seems counterintuitive, but could be explained by the fact that L. paradoxa possesses two pairs of internal connected nares (nares perforatae; Hyrtl, 1843), that are clearly visible during a ventilatory cycle (Goeldi, 1898). The outgoing airflow could be guided by a jet stream from the lungs to the posterior pair of internal nares, assisted by a series of longitudinal folds along the dorsal part of the mouth (Fig. 3). The expiratory flow leaving the animal through the nares could therefore be separated from the inspiratory flow occurring through the mouth, and the negative pressure measured could be explained by the expansion of the buccal cavity. As shown by Brainerd (1994), buccal diameter increases in L. paradoxa concomitantly with a decrease in lung diameter, and even after the lung has reached its minimum diameter of 40%, the buccal cavity diameter continues to increase. Once lung diameter stops decreasing, buccal cavity still increases some 30% in diameter (Brainerd, 1994). These findings suggest that expiratory airflow must occur through an expanding buccal cavity, resulting in the mixing of fresh and exhaled air. Our recordings suggest that the constant increase in buccal volume with the accompanying outflow of air, maintains pressure in the buccal cavity constantly sub-atmospheric. At point ‘c’ in Fig. 1, buccal pressure starts to rise sharply, becoming positive, but expiratory airflow continues. This suggests that at the first compression of the buccal cavity, some air is being expelled through the mouth, whereas the remaining air is being pumped into the lungs. The following expansions and compressions of the buccal cavity result in buccal cavity pressure oscillating between increasingly positive and small negative pressures. The increase in buccal pressure throughout the sequence of lung inflations could be explained in that as the maximal volume is approached, the total compliance of the system decreases. The buccal pressure oscillations observed at the end of the ventilatory cycle may be associated to the expulsion of air from the buccal cavity through the opercular openings (G.S.F·S, personnel observation; Goeldi, 1898; Bishop and Foxon, 1968), once the lungs have been filled completely. Ventilation data obtained in the present study are comparable to results obtained in previous studies using L. paradoxa at 25 °C (Table 3) and further confirm that V̇ E is being adjusted by changing fR, since expiratory VT is typically maintained at 25.0 ± 4.6 mL BTPS kg−1. 163
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ventilation, the buccal PCO2 is nearly zero; then we would expect to find PBCO2 during a ventilatory cycle (end-tidal CO2) to be much lower than described in the present study (~ 19 mmHg). Rather, we find a very small lung-buccal PCO2 difference (Table 1), which supports the idea of a near complete replacement of lung air during one ventilatory cycle. These data are in agreement with Fernandes et al. (2005) for the toads. In conclusion, L. paradoxa is able to renew almost all of its pulmonary air during one ventilatory cycle, releasing most of the air to the outside during the expansion of the buccal cavity. The degree of mixing within the buccal cavity is very small, as evidenced by the small dead space calculated, suggesting the presence of a ‘jet stream’ of used air leaving the lungs and rapidly passing through a slowly expanding buccal cavity. A jet stream mechanism may have evolved independently in lungfish and some anuran amphibians. But if considered being a plesiomorphic breathing mechanism of buccal pump breathing sarcopterygians, jet streaming would have significant implications on the evolution of air breathing in early tetrapods.
responses to aerial hypoxia and temperature in the south American lungfish Lepidosiren paradoxa. J. Therm. Biol. 36, 521–526. da Silva, G.S.F., Ventura, D.A.D.N., Zena, L.A., Giusti, H., Glass, M.L., Klein, W., 2017. Effects of aerial hypoxia and temperature on pulmonary breathing pattern and gas exchange in the south American lungfish, Lepidosiren paradoxa. Comp. Biochem. Physiol. A. 207, 107–115. De Jongh, H.J., Gans, C., 1969. On the mechanism of respiration in the bullfrog Rana catesbeiana. A reassessment. J. Morphol. 127, 259–290. de Moraes, M.F.P.G., Höller, S., da Costa, O.T.F., Glass, M.L., Fernandes, M.N., Perry, S.F., 2005. Morphometric comparison of the respiratory organs in the south American lungfish Lepidosiren paradoxa (Dipnoi). Physiol. Biochem. Zool. 78, 546–559. Dean, B., 1906. Notes on the living specimens of the Australian lungfish, ceratodus forsteri, in the zoological Society's collection. Proc. Zool. Soc. Lond. 76, 168–178. Dejours, P., 1981. Principles of Comparative Respiratory Physiology, 2nd ed. Elsevier/ North-Holand, Amsterdam. DeLaney, R.G., Laurent, P., Galante, R., Pack, A.I., Fishman, A.P., 1983. Pulmonary mechanoreceptors in the Dipnoi lungfish Protopterus and Lepidosiren. Am. J. Phys. 244, R418–R428. Fernandes, M.S., Giusti, H., Glass, M.L., 2005. An assessment of dead space in pulmonary ventilation of the toad Bufo schneideri. Comp. Biochem. Physiol. A 142, 446–450. Fritsche, R., Axelsson, M., Franklin, C.E., Grigg, G.G., Holmgren, S., Nilsson, S., 1993. Respiratory and cardiovascular responses hypoxia in the Australian lungfish. Respir. Physiol. 94, 173–187. Gans, C., De Jongh, H.J., Farber, J., 1969. Bullfrog (Rana catesbeiana) ventilation: how does the frog breathe? Science. 163, 1223–1225. Glass, M.L., 2010. Respiratory function in lungfish (Dipnoi) and a comparison to land vertebrates. In: Jorgensen, J.M., Joss, J. (Eds.), The Biology of Lungfishes. Taylor & Francis, pp. 265–282. Glass, M.L., Boutilier, R.G., Heisler, N., 1983. Ventilatory control of arterial PO2 in the turtle, Chrysemys picta bellii: effects of temperature and hypoxia. J. Comp. Physiol. B. 151, 145–153. Goeldi, E.A., 1898. On the Lepidosiren of the amazons; being notes on five specimens obtained between 1895-97, and remarks upon an example living in the Para museum. Trans. zool. Soc. Lond. 14, 413–420. Graham, J.B., 1997. Air-Breathing Fishes. Evolution, Diversity, and Adaptation. Academic Press. Grigg, G.C., 1965. Studies on the Queensland Lungfish, Neoceratodus forsteri (Krefft) 1. Anatomy, histology, and functioning of the lung. Austral. J. Zool. 13, 243–254. Günther, C.A.I.G., 1871. Description of ceratodus, a genus of gadoid fishes, recently discovered in rivers of Queensland. Australia. Trans. Roy. Soc. Lond. 161, 511–792. Harder, V., 2001. Untersuchungen zur bimodalen Atmung des südamerikanischen Lungenfisches Lepidosiren paradoxa (Fitz.) - in situ und in vivo. (Dissertation). Universität Düsseldorf. Shaker-Verlag, Aachen. Hughes, G.M., Weibel, E.R., 1976. Morphometry of fish lungs. In: Hughes, G.M. (Ed.)Respiration of Amphibious Vertebrates. Academic Press, London, pp. 213–232. Hyrtl, J., 1843. Lepidosiren paradoxa. Monographie. Abh. Königl. Böhm. Ges. Wiss. 5, 606–668. Johansen, K., Lenfant, C., 1967. Respiratory function in the south American lungfish, Lepidosiren paradoxa. J. Exp. Biol. 46, 205–218. Johansen, K., Lenfant, C., 1968. Respiration in the African lungfish Protopterus aethiopicus: II control of breathing. J. Exp. Biol. 49, 453–468. Klika, E., Lelek, A., 1967. A contribution to the study of the lungs of the Protopterus annectens and Polypterus senegalensis. Folia Morphol. (Warsz) 15, 168–175. Lenfant, C., Johansen, K., Handson, D., 1970. Bimodal gas exchange and ventilationperfusion relationship in lower vertebrates. Fed. Proc. 29, 1124–1129. Maina, J.N., Maloiy, G.M.O., 1985. The morphometry of the lung of the African lungfish (Protopterus aethiopicus): its structuralfunctional correlations. Proc. R. Soc. London 224B, 399–420. McMahon, B.R., 1969. A functional analysis of the aquatic and aerial respiratory movements of an african lungfish, Protopterus aethiopicus, with reference to the evolution of the lung ventilation mechanism in vertebrates. J. Exp. Biol. 51, 407–430. McMahon, B.R., 1970. The relative efficiency of gaseous exchange across the lungs and gills og an African lungfish Protopterus aethiopicus. J. Exp. Biol. 21, 1–15. Owen, R., 1841. Description of the Lepidosiren annectens. Trans. Linn. Soc., London 18, 327–361. Parker, W.N., 1892. On the anatomy and physiology of Protopterus annectens. Trans. R. Irish Acad. 30, 109–230. Poll, M., 1962. Étude sur la structure adulte et la formation des sacs pulmonaires des Protoptères. Ann. Mus. R. l'Afrique Cent. Sci. Zool. 108, 130–172. Quekett, J., 1844. On a peculiar arrangement of blood-vessels in the air-bladder of fishes, with some remarks on the evidence which they afford of the true function of that organ. Trans. R. Micro. Soc., London 1, 99–108. Sanchez, A.P., Glass, M.L., 2001. Effects of environmental hypercapnia on pulmonary ventilation of the South American lungfish. J. Fish Biol. 58, 1181–1189. Sanchez, A.P., Hoffman, A., Rantin, F.T., Glass, M.L., 2001b. The relationship between pH of the cerebro-spinal fluid and pulmonary ventilation of the south American lungfish, Lepidosiren paradoxa. J. Exp. Zool. 290, 421–425. Sanchez, A.P., Soncini, R., Wang, T., Koldkjær, P., Taylor, E.W., Glass, M.L., 2001c. The differential cardio-respiratory responses to ambient hypoxia and systemic hypoxaemia in the south American lungfish, Lepidosiren paradoxa. Comp. Biochem. Physiol. 130, 677–687. Sanchez, A.P., Giusti, H., Bassi, M., Glass, M.L., 2005. Acid–base regulation in the south American lungfish, Lepidosiren paradoxa: effects of prolonged hypercarbia on blood gases and pulmonary ventilation. Physiol. Biochem. Zool. 78, 908–915. Vitalis, T.Z., Shelton, G., 1990. Breathing in Rana pipiens: the mechanism of ventilation. J. Exp. Biol. 154, 537–556.
Conflict of interests The authors confirm that there is no conflict of interest Acknowledgements Funding: This study was financially supported by the Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq), through the National Institute of Science and Technology on Comparative Physiology (INCT em Fisiologia Comparada, proceeding nos. 2008/ 57712-4 and 573921/2008-3). G.S.F. da Silva was supported by a Young Investigator Award (FAPESP no 2013/17606-9 and 2014/ 12190-1) and is currently supported by CNPq (403769/2016-7). W. J. Minto was the recipient of a FAPESP postdoc fellowship (no. 2009/ 53916-7). References Amin-Naves, J., Giusti, H., Glass, M.L., 2004. Effects of acute temperature changes on aerial and aquatic gas exchange, pulmonary ventilation and blood gas status in the south American lungfish, Lepidosiren paradoxa. Comp. Biochem. Physiol A 138, 133–139. Amin-Naves, J., Giusti, H., Hoffmann, A., Glass, M.L., 2007a. Components to the acid–base related ventilatory drives in the south American lungfish Lepidosiren paradoxa. Respir. Physiol. Neurobiol. 155, 35–40. Amin-Naves, J., Giusti, H., Hoffmann, A., Glass, M.L., 2007b. Central ventilatory control in the south American lungfish, Lepidosiren paradoxa: distinct responses to pH and CO2. J. Comp. Physiol. B. 177, 529–534. Bassi, M., Klein, W., Fernandes, M.N., Perry, S.F., Glass, M.L., 2005. Pulmonary oxygen diffusing capacity of the south American lungfish Lepidosiren paradoxa: physiological values by the Bohr integration method. Physiol. Biochem. Zool. 78, 560–569. Bassi, M., Giusti, H., da Silva, G.S., Amin-Naves, J., Glass, M.L., 2010. Blood gases and cardiovascular shunt in the south American lungfish (Lepidosiren paradoxa) during normoxia and hyperoxia. Respir. Physiol. Neurobiol. 173, 47–50. Bishop, I.R., Foxon, G.E.H., 1968. The mechanism of breathing in the south American lungfish, Lepidosiren paradoxa; a radiological study. J. Zool. Lond. 154, 263–271. Brainerd, E.L., 1994. The evolution of lung gill bimodal breathing and the homology of vertebrate respiratory pumps. Amer. Zool. 34, 289–299. Brainerd, E.L., Ferry-Graham, L.A., 2006. Mechanics of respiratory pumps. In: Shadwick, R.E., Lauder, G.V. (Eds.), Fish Biomechanics Fish Physiology series. 23. pp. 1–28. Brainerd, E.L., Ditelberg, J.S., Brambçe, D.M., 1993. Lung ventilation in salamanders and the evolution of vertebrate air-breathing mechanisms. Biol. J. Linn. Soc. 49, 163–183. Brett, S.S., Shelton, G., 1979. Ventilatory mechanisms of the amphibian, Xernopus laevis; the role of the buccal force pump. J. Exp. Biol. 80, 251–269. Brinkmann, H., Venkatesh, B., Brenner, S., Meyer, A., 2004. Nuclear protein-coding genes support lungfish and not the coelacanth as the closest living relatives of land vertebrates. Proc. Natl. Acad. Sci. 101, 4900–4905. Clement, A.M., Long, J.A., 2010. Air-breathing adaptation in a marine Devonian lungfish. Biol. Lett. 6, 509–512. da Silva, G.S.F., Giusti, H., Sanchez, A.P., do Carmo, J.M., Glass, M.L., 2008. Aestivation in the south American lungfish, Lepidosiren paradoxa: effects on cardiovascular function, blood gases, osmolality and leptin levels. Respir. Physiol. Neurobiol. 164, 380–385. da Silva, G.S.F., Giusti, H., Branco, L.G.S., Glass, M.L., 2011. Combined ventilatory
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602–609. Zardoya, R., Cao, Y., Hasegawa, M., Meyer, A., 1998. Searching for the closest living relative(s) of tetrapods through evolutionary analyses of mitochondrial and nuclear data. Mol. Biol. Evol. 15, 506–517. Zena, L.A., Bícego, K.C., da Silva, G.S.F., Giusti, H., Glass, M.L., Sanchez, A.P., 2017. Acute effects of temperature and hypercarbia on cutaneous and branchial gas exchange in the south American lungfish, Lepidosiren paradoxa. J. Therm. Biol. 63, 112–118.
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This article is part of a Special issue on Physiology from the Neotropics
Buccal jet streaming and dead space determination in the South American lungfish, Lepidosiren paradoxa☆ Walter J. Mintoa, Humberto Giustia, Mogens L. Glassa,1, Wilfried Kleinb,c, Glauber S.F. da Silvac,d,
T ⁎
a
Faculty of Medicine of Ribeirão Preto, University of São Paulo, Ribeirao Preto, SP, Brazil School of Philosophy, Sciences and Literature of Ribeirão Preto, University of São Paulo, Ribeirão Preto, SP, Brazil c National Institute of Science and Technology on Comparative Physiology, Rio Claro, SP, Brazil d Institute of Biological Science, Department of Physiology and Biophysics, Federal University of Minas Gerais, Belo Horizonte, MG, Brazil b
A R T I C LE I N FO
A B S T R A C T
Keywords: Buccal pump Air-breathing Lung gas End-tidal CO2 Effective ventilation Bohr equation Lungfish
The “jet stream” model predicts an expired flow within the dorsal part of the buccal cavity with small air mixing during buccal pump ventilation, and has been suggested for some anuran amphibians but no other species of air breathing animal using a buccal force pump has been investigated. The presence of a two-stroke buccal pump in lungfish, i.e. expiration followed by inspiration, was described previously, but no quantitative data are available for the dead-space of their respiratory system and neither a detailed description of airflow throughout a breathing cycle. The present study aimed to assess the degree of mixing of fresh air and expired gas during the breathing cycle of Lepidosiren paradoxa and to verify the possible presence of a jet stream during expiration in this species. To do so, simultaneous measurements of buccal pressure and ventilatory airflows were carried out. Buccal and lung gases (PCO2 and PO2) were also measured. The effective ventilation was calculated and the dead space estimated using Bohr equations. The results confirmed that the two-stroke buccal pump is present in lungfish, as it is in anuran amphibians. The present approaches were coherent with a small dead space, with a very small buccal-lung PCO2 difference. In the South American lungfish the dead space (VD) as a percentage of tidal volume (VT) (VD / VT) ranged from 4.1 to 12.5%. Our data support the presence of a jet stream and indicate a small degree of air mixing in the buccal cavity. Comparisons with the literature indicate that these data are similar to previous data reported for the toad Rhinella schneideri.
1. Introduction The six species of extant Dipnoi (Neoceratodus forsteri; Lepidosiren paradoxa; Protopterus aethiopicus, P. amphibius, P. annectens, P. dolloi) possess lungs as their principal air-breathing organs (Graham, 1997). The morphological characteristics of dipnoan lungs have been studied for centuries (Owen, 1841; Quekett, 1844; Hyrtl, 1843; Günther, 1871; Parker, 1892; Poll, 1962; Grigg, 1965; Klika and Lelek, 1967; Hughes and Weibel, 1976; Maina and Maloiy, 1985; de Moraes et al., 2005). Several aspects of lung ventilation, gas exchange, blood gas chemistry, or chemoreception have also been of interest for comparative physiologists, especially over the last 50 years (e.g. Johansen and Lenfant, 1967; Johansen and Lenfant, 1968; Lenfant et al., 1970; McMahon, 1970; Fritsche et al., 1993; Amin-Naves et al., 2004; DeLaney et al., 1983; Sanchez and Glass, 2001; Glass, 2010; Sanchez and Glass, 2001;
Sanchez et al., 2001b; Bassi et al., 2005; Bassi et al., 2010; Sanchez et al., 2005; Amin-Naves et al., 2007a, 2007b; da Silva et al., 2008, 2017; Zena et al., 2017). Such attention given to morphology and physiology of lungfish can be explained by their phylogenetic position close to the origin of the Sarcopterygii (Brinkmann et al., 2004; Yokobori et al., 1994; Zardoya et al., 1998) and their ability to exchange gases bimodally with water (across gills and/or integument) and air (using lungs), representing the remaining living species of a lineage of vertebrates that evolved air breathing in the mid-Devonian (~380 Ma; Clement and Long, 2010). One aspect of lungfish respiration, however, has not received much attention up to now: the breathing mechanism (Bishop and Foxon, 1968; McMahon, 1969; Brainerd, 1994). While studies of the buccal pump mechanism have been carried out on L. paradoxa (Bishop and Foxon, 1968; Brainerd, 1994) and P. aethiopicus (McMahon, 1969;
☆
This article is part of a special issue entitled: Physiology from the Neotropics, edited by: Dr. Kenia Bicego, Dr. Luciane Gargaglioni and Dr. Mike Hedrick Corresponding author at: Departamento de Fisiologia e Biofísica, Instituto de Ciências Biológicas; Universidade Federal de Minas Gerais, Av. Presidente Antonio Carlos, 6627. ICB/UFMG, Depto Fisiologia e Biofísica. Sala 158, Bloco B4-158, Pampulha, 31270-901 Belo Horizonte, MG, Brazil. E-mail address: [email protected] (G.S.F. da Silva). 1 In memorian, Deceased October 4th, 2018. ⁎
https://doi.org/10.1016/j.cbpa.2019.05.026 Received 3 March 2019; Received in revised form 8 May 2019; Accepted 20 May 2019 Available online 10 June 2019 1095-6433/ © 2019 Elsevier Inc. All rights reserved.
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2. Material and methods
Brainerd et al., 1993), only superficial descriptions of breathing behavior are available for N. forsteri (Dean, 1906; Grigg, 1965). It seems clear, however, that all lungfish use a two-stroke buccal pump, composed of one expiration and at least one inspiration, to renew the air within the lungs (Brainerd, 1994). The events during one breathing cycle can be described as follows (Brainerd and Ferry-Graham, 2006): 1) Air breathing starts with the animal putting its head out of the water and the mouth being opened; 2) The buccal cavity expands to draw in oxygen-rich air; 3) As the buccal cavity expands, the glottis opens and the lungs empty through elastic recoil of body wall and lungs; 4) Once the buccal cavity has been fully expanded, the mouth closes and the air trapped within the buccal cavity is pushed into the lungs. Since fresh air and gas expired from the lungs enter the buccal cavity during buccal expansion, some degree of mixing of oxygen-rich and oxygen-poor gas occurs during the first phase of the two-stroke buccal pump, reducing the amount of oxygen entering the lungs. McMahon (1969) estimates the gas mixing to be 20–40% of expired air being returned to the lungs in P. aethiopicus. A two-stroke buccal pump ventilation has also been identified in anuran amphibians. De Jongh and Gans (1969) and Gans et al. (1969) performed classical experiments using the bullfrog (Lithobates catesbeianus) to describe the complex functioning of the buccal pump. De Jongh and Gans (1969) identified three different types of cyclical phenomena associated to buccal movements. 1) Oscillations of the buccal floor to ventilate the buccal cavity through the open nares. 2) Ventilatory cycles to renew the air within the lungs through coordinated opening and closing of nares and glottis. 3) Inspiratory movements with pulmonary ventilation interspaced by non-ventilatory periods. One fundamental proposal of de Jongh and Gans (1969) was the presence of an expiratory airflow, known as jet stream, postulated to result in minimal mixing of oxygen rich air in the buccal cavity with the oxygen poor air leaving the lungs during expiration, and passing rapidly along the dorsal part of the buccal cavity. For Lithobates pipiens a similar ventilatory pattern as in the bullfrog has been described, but no evidence for the presence of an expiratory jet stream has been found (Vitalis and Shelton, 1990), since the air being pumped into the lungs during inspiration represents a mixture of air, containing 30–50% of air from the previous expiration (West and Jones, 1975). Brett and Shelton (1979), studying Xenopus laevis, found in this species a ventilatory cycle starting with an expiration that is followed by an inspiration, thereby suggesting no mixing of fresh and used air within the buccal cavity. Fernandes et al. (2005), investigating the ventilatory pattern and jet stream hypothesis in the toad Rhinella schneideri, found a buccal cavity dead-space representing 30–40% of tidal volume, suggesting some degree of inspiratory and expiratory flow separation, with limited mixing within the buccal cavity. Regarding the ventilatory cycle of L. paradoxa, it has to be assumed that this species possibly expires almost all of the air contained within the lungs at the beginning of a ventilatory cycle. This was demonstrated by Bishop and Foxon (1968) using X-ray imaging of a breathing cycle, where the lungs were no longer visible during expiration due to a nearcomplete loss of air, also has been suggested by da Silva et al. (2017), based on measurements of expiratory tidal volume. Since pulmonary dead space seems to be very small, a significant amount of gas mixing may only be possible during the expansion phase of the two-stroke buccal pump, when air is being expired from the lungs and fresh air is being drawn into the buccal cavity. To determine the degree of mixing of fresh air and expired gas during the breathing cycle of L. paradoxa and to verify the possible presence of a jet stream during expiration in this species, we performed measurements of buccal pressure, buccal and pulmonary PCO2 and PO2, pulmonary ventilation and calculated dead space and effective ventilation by using the Bohr dead space equation (see Eq. 3).
2.1. Animals The lungfish specimens of Lepidosiren paradoxa were collected in the region of Cuiabá – MT (Brazil) and transported to the University of São Paulo, campus Ribeirão Preto (FMRP/USP) (authorized by IBAMA # 02027. 002172/2005–68). Upon arrival, the animals were placed in tanks of 1000 L with aerated and temperature-controlled water (25 °C) and were fed three times a week with chicken liver. Before the experiments, each animal remained 24 h without access to food. The experimental procedures were approved by the local ethics and animal care committee (Proc. # 0762005). Ten animals (N = 10) of both sexes were used in the present study. In order to avoid running experiments during the aestivation period (dry season – from May to September) all procedures took place during animal's active season, from October (2011) to April (2012).
2.2. Surgical procedures Animals were anesthetized by immersion in a solution of benzocaine to 1:1 g/L as extensively described in our previous studies (Amin-Naves et al., 2007a, 2007b; da Silva et al., 2011; da Silva et al., 2008). Under anesthesia, lungs and buccal cavity were cannulated in different groups of animals. For cannulation of the lung, anesthetized animals were transferred to the surgical apparatus, in which anesthesia was maintained by switching to a diluted (1:4) solution of benzocaine with water through a catheter onto the gills. An incision (~2 cm) was made into the body wall in the dorsal caudal region to expose the posterior part of the lungs. Care was taken not to damage any closely positioned vessels. Then, a cannula (PE 50 connected to a PE 90) was inserted into the lung through a small incision and secured externally around the lung. The catheter was exteriorized and the skin sutured. The buccal cavity was cannulated using a polyethylene cannula (PE 50) connected to a PE 90, which was inserted into the buccal cavity through a micro incision in the lateral buccal floor. This cannula was used to collect air samples (for PO2 and PCO2 measurements) and to record pressures. After surgical procedures, animals were allowed to recover for at least one day before starting the experimental protocols.
2.3. Measurements of pulmonary ventilation The experimental apparatus used for pulmonary ventilation recordings was the same as described in our previous studies (da Silva et al., 2011, 2017). Briefly, animals were placed into a closed plastic box that contained perforated sidewalls for water circulation. An inverted funnel was attached to the lid of the plastic box, representing the only place for the animal to surface to breathe air. A homemade pneumotachograph (formed by a set of parallel tubes) was attached to the funnel to record ventilatory airflow. This setup was completely immersed into a much larger Styrofoam box and a submerged pump, which was placed outside of the plastic box, was used to maintain water circulation within the experimental apparatus. Animals were placed into this apparatus for a period of 12 h for acclimatization before starting the experiment. Pulmonary ventilation was measured using the pneumotachographic method described by Glass et al. (1983) for aquatic animals. A differential pressure transducer (141 ML spirometer; ADInstruments, Sydney, Australia) was connected to the pneumotach and recorded breathing-related pressure oscillations. The system was calibrated by injecting and withdrawing a known volume of air (10 mL). This method was used for determining air-breathing frequency (fR) and tidal volume (VT, mL kg−1). The calculation of total lung ventilation (VĖ TOT, mL BTPS Kg−1 h−1) was obtained by multiplying fR by VT. 160
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as a percentage of tidal volume (VT) (Fernandes et al., 2005):
2.4. Experimental protocols
VD/ VT = (PL CO2 –PB CO2)/ PL CO2
All the experiments were carried out during the active season, daytime at normoxic (20.95% O2) and normocarbic (0.03% CO2) conditions (at both gas and water phase) and at controlled temperature of 25 °C.
(3)
where: PBCO2 represents the CO2 partial pressure in the buccal cavity during a breathing episode. This calculation is usually based on intrapulmonary PCO2 (PLCO2) measurements compared to end-tidal PCO2 (PETCO2) during expiration of the breathing episode (Fernandes et al., 2005). However, it is not possible to place a mask onto the lungfish in order to measure end-tidal gas compositions, thus we used PBCO2 as substitute for PETCO2. Similar to alveolar ventilation (V̇ A), the effective pulmonary ventilation (V̇ EF) was obtained as (Wang et al., 1998; Fernandes et al., 2005):
2.4.1. Measurements of buccal pressure To measure variations in buccal pressure associated to the ventilatory cycle, an animal was placed into the experimental apparatus and had the buccal catheter connected to a pressure transducer (MLT0380; ADInstruments, Sydney, Australia). The transducer signals passed through a single-channel Bridge Amp (FE221, ADInstruments) and were recorded using a data acquisition system (PowerLab 4/30; AdInstruments) and analyzed using LabChart 7 software (AdInstruments).
V̇ Eff = (V̇ CO2•RT )/P LCO2
(4)
where: V̇ Eff gives the effective ventilation of the lung, V̇ CO2 the CO2 elimination through the lung, PLCO2 the intrapulmonary (lung) PCO2, R the gas constant, and T the absolute temperature (K). V̇ CO2 was not measured in the present study and hence the value used to calculate V̇ Eff was obtained from our previous article (da Silva et al., 2017). Knowing V̇ Eff and V̇ ETot, dead-space was also calculated as:
2.4.2. Measurements of partial pressure of CO2 (PCO2) and O2 (PO2) in lung and buccal compartments The purpose of this protocol was to obtain the partial pressure of CO2 and O2 within the buccal cavity and lungs of unanaesthetized L. paradoxa. Animals within the experimental apparatus had the pulmonary cannula connected to a 5 mL syringe. Firstly, to avoid any collapsed areas within the lungs, a volume of 5 mL of room air was injected. Then, the same volume of air was withdrawn from the lung after four different time-points (1, 10, 20 and 40 min). The lung air samples were injected into a gas analyzer (ML206 ADInstruments, Sydney, Australia) and the gases were recorded using the same data acquisition system as for the pressure measurements. Importantly, the syringe was sealed using a three-way stopcock to avoid any leaking and minimize any possible mixture with room air. Air samples from previously mentioned time points were averaged in each animal and considered for the further calculations, where values of lung PO2 and PCO2 (PLO2 and PLCO2, respectively) were needed. The gas composition within the buccal cavity was measured through the buccal catheter being connected to the gas analyzer and data acquisition system (ADInstruments). A small sampling flow rate was obtained by the analyzer built-in pump. The oscillations in O2 and CO2 were monitored to determine gas partial pressures in the buccal cavity (PBO2 and PBCO2) during spontaneous air breathing (ventilatory movements). To accomplish these measurements, the animal was kept not totally submerged in water, but rather, in an aquarium with a thin layer of water, enough to keep it humidified. This procedure allowed us to perform buccal gas sampling without any water from the buccal cavity being sucked to the gas analyzer. Gas recordings were obtained as percentage (%) or fraction and were subsequently converted to mmHg considering a barometric pressure of 710 mmHg and the partial pressure of water vapor at 25 °C.
̇ –VEff ̇ )/ fR VD = (VETot
(5)
Knowing tidal volume (VT), the relationship VD / VT was obtained. Lung and buccal PO2 and PCO2 were compared using Student's ttest. All data are reported as mean and standard error (mean ± SE). Significance level was P < .05. The N-value was 5 for each data set. 4. Results Fig. 1 depicts a representative buccal pressure profile along with airflow measurements during a breathing event in the lungfish. A
3. Equations used and statistics The lung O2 extraction coefficient (EO2) (which expresses the percentage of O2 absorbed by the pulmonary capillaries) and the respiratory exchange ratio (RER) were calculated following (Dejours (1981):
E O2 (%) = [PI O2 − PL O2)/ PI O2]•100
(1)
RER = PL CO2 /(PI O2 − PL O2)
(2)
Fig. 1. Measurements of buccal pressure (A) and ventilatory air flow (B) in L. paradoxa at 25 °C. The letters ‘a’ to ‘k’ represent events associated to changes in buccal pressure and airflow. ‘a’ represents the start of a ventilatory cycle followed by a decrease in buccal pressure to sub-atmospheric values; ‘b’ marks initiation of expiratory flow; ‘c’ represents the increase in buccal pressure associated with air being pushed from the buccal cavity into the lungs; ‘d’, ‘f’, ‘h’ give peak buccal pressures and mark initiation of inspiratory airflow; ‘e’, ‘g’, ‘i’ represent inspiratory airflow and the initial increase of buccal pressure associated with air being pushed into the lungs; ‘j’ marks the last peak buccal pressure and flow measurements returning to zero flow; ‘k’ marks buccal pressure oscillations associated to the elimination of remaining air from the buccal cavity.
where: PIO2 gives the oxygen partial pressure of inspired air (in mmHg), PLO2 the oxygen partial pressure in the lungs (in mmHg), and PLCO2 the lung carbon dioxide partial pressure (mmHg). Eq. 2 is a simple rearrangement of the alveolar gas equation, where alveolar PO2 and PCO2 (PAO2 and PACO2) were replaced by the measured values of PLO2 and PLCO2. The Bohr dead-space equation was used to estimate dead space (VD) 161
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Table 1 Respiratory variables measured and calculated in the lungfish L. paradoxa in the present study. Respiratory variables
Mean ± SE
V̇ E (mL BTPS h−1 kg−1) Effective V̇ Eff (mL BTPS h−1 kg −1) VT (mL kg−1) fR (breaths h−1) Lung PCO2 (mmHg) Buccal PCO2 (mmHg) Lung PO2 (mmHg) Buccal PO2 (mmHg) RER Extraction Coefficient (%) EO2 (%) = [PIO2-PLO2)/PIO2] •100
89.85 ± 9.70 82.77 ± 5.80 27.08 ± 2.80 2.0 ± 0.2 20.63 ± 1.60 19.90 ± 2.40 93.80 ± 6.80 87.40 ± 11.10 0.42 ± 0.04 35.3 ± 4.7
PCO2 and PO2 values were statistically not different in the lung and buccal compartments (PCO2; buccal vs lung: p = .8 and PO2; buccal vs lung: p = .64). Table 2 Calculation of dead-space in percentage of tidal volume (VD/VT) and comparison between the South American lungfish L. paradoxa and the Toad Rhinella schneideri.
Fig. 2. Values of gases partial pressures in the lung (circles) and buccal (squares) compartments. PCO2 values are depicted in panel A and PO2 in the panel B. Each point represents the value of an individual animal, while horizontal lines (middle and top-bottom) represent mean and standard error. There is no statistical difference in the PCO2 and PO2 between lung and buccal compartments (N = 5).
1) VD = (VTOT – VEff) / fR
Lungfish
Toad
Effective ventilation (mL BTPS •min−1 kg −1) Total ventilation (mL BTPS •min−1 kg −1) Breathing frequency (cycles •min−1) Tidal volume (VT) (mL •kg−1) Dead space volume (VD) (mL •kg−1) (VD/VT) 100%
1.37 1.48 0.03 27.0 3.54 12.5
7.00 12.10 5.80 2.4 0.90 37.0
2) VD/VT = (PLCO2 – PBCO2) / PLCO2 PLCO2 (mmHg) PBCO2 (mmHg) (VD/VT) 100%
20.6 19.9 4.12
10.9 7.6 30.2
3) Substitution of PLCO2 by PaCO2 PaCO2 (mmHg)⁎ (VD/VT) 100%
21.8 8.7
12.3 49.9
⁎ PaCO2 value has been calculated as average from several studies on the lungfish L. paradoxa at 25 °C in air and water normoxia (for details see review article submitted). For purposes of comparison, the numbers presented for both, lungfish (present study) and toad (Fernandes et al., 2005) are mean values.
typical ventilatory cycle in L. paradoxa was characterized by a slight increase in buccal pressure (‘a’ in Fig. 1) followed by a decrease in buccal pressure to sub-atmospheric values. Once negative pressure stopped decreasing, expiratory flow initiated (‘b’) and continued for about 4 s. With expiratory flow still occurring, buccal pressure increased sharply to positive values (‘c’). After reaching a peak buccal pressure of about 6 mmHg (‘d’), pressure in the buccal cavity fell to negative values again, coinciding with an inspiratory flow. Buccal pressure rose again sharply for another three times to positive values (9–12 mmHg), always accompanied by a decreasing respiratory flow (‘e’, ‘g’, ‘i’). Following each of these increasing buccal pressures were three sharply decreasing buccal pressures (‘f’, ‘h’, ‘j’), that were coincident with inspiratory air flows, with the exception of the last decrease in buccal pressure (‘j’). At this moment (‘j), ventilatory flow measurements returned to zero flow and buccal pressure oscillated between positive and negative values before returning to zero. All compressions and expansions of the buccal cavity (from ‘c’ to ‘k’) took 0.8 to 1.0 s each. Partial pressures of O2 and CO2 from the lungs and the buccal cavity are given in Fig. 2 and Table 1 and the mean values of V̇ E, VT and fR, along with other measured and calculated respiratory variables are presented in the Table 1 (all under normoxic conditions and at 25 °C). The PaCO2 and V̇ CO2 values were taken from previous studies (review article; da Silva et al., 2017) and used to calculate effective ventilation (eq. 4), which corresponded to a large fraction of the total ventilation. This result agrees with the relatively small dead-space (VD) found. Although lung PO2 and PCO2 tended to be greater than values measured in the buccal cavity, no statistical differences were found between lung and buccal gas composition (Fig. 2). The O2 extraction
coefficient, calculated using the inspired-lung PO2 difference (eq. 1), was found to be 35.3 ± 4.7%. Table 2 demonstrates the calculations of dead-space in percentage of tidal volume (VD / VT) and comparisons between the South American lungfish L. paradoxa and the toad Rhinella schneideri as described by Fernandes and et al. (2005). Using the total ventilation, effective ventilation and breathing frequency, the VD / VT ratio was estimated to be 12.5% (eq. 5). In absolute values, the deadspace was calculated and estimated to be 3.5 mL kg−1. Based on the small difference in lung and buccal PCO2, the dead-space calculated using the Bohr equation (eq. 3) corresponded to 4.1 ± 6% of the tidal volume. Substitution of lung PCO2 by PaCO2 gave a dead-space of 8.7% of tidal volume. Thus, in the South American lungfish the VD / VT ranged from 4.1 to 12.5%, which were smaller than those values reported for R. schneideri. 5. Discussion Pulmonary airflow measurements in the current study confirm the presence of a two-stroke breathing mechanism in L. paradoxa, composed of a single expiration followed by 2–4 buccal inspirations. Our recordings of buccal cavity pressure, however, were different from previous recordings, since a negative buccal pressure was found at the beginning of each ventilatory cycle, always coinciding with expiratory flow. While McMahon (1969) and Brainerd et al. (1993) did record in P. aethiopicus small negative pressures in the buccal cavity at the 162
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Table 3 Mean values reported in the literature for pulmonary ventilation (V̇ E, ml BTPS kg−1 h−1), tidal volume (VT, ml BTPS kg−1), and breathing frequency (fR, breaths h−1) in L. paradoxa at 25 °C.
Fig. 3. Postulated airflow pattern during the first expansion of the buccal cavity in Lepidosiren paradoxa in frontal view of the open mouth (A) and in lateral view (B). Blue traces in A show the postulated jet stream of air along the dorsal part of the buccal cavity, being guided by dorsal folds towards the posterior internal nares and exiting through the anterior internal nares. Blue arrows in B show expiratory airflow leaving through the anterior internal nares, while the red arrow represents inspiratory flow. Panel A modified from Hyrtl (1843) and panel B modified from Goeldi (1898).
Reference
V̇ E
VT
fR
Amin-Naves et al. (2004) Amin-Naves et al. (2007a) Amin-Naves et al. (2007b) Bassi et al. (2010) da Silva et al. (2011) da Silva et al. (2017) Harder (2001) Sanchez et al. (2001a) Sanchez et al. (2001c) Sanchez et al. (2005) Present study
162.0 22.1 39.0 63.7 64.9 73.4 42.0 88.2 103.0 43.0 89.9
29.0 15.0 20.0 20.5 29.8 27.3 27.8 24.0 26.0 28.0 27.1
5.4 1.0 1.3 3.2 2.1 2.6 1.4 5.5 4.0 1.7 2.0
Similarly, the data obtained regarding lung PCO2 and PO2 are in the same range as in previous studies (Johansen and Lenfant, 1968; Bassi et al., 2005; da Silva et al., 2011). Fernandes et al. (2005), using identical methods to calculate ventilatory dead space in Rhinella schneideri, found dead space to make up between 30 and 50% of VT. No other estimates for ventilatory dead space are available for amphibians, nor any air-breathing fish. A comparison with amniotes seems not valid, since their respiratory system is composed of lungs being connected through a trachea to the mouth cavity, a morphological feature not seen in any air-breathing fish and amphibian. In lungfish, the lungs are connected through a short duct to the esophagus, and this connecting duct is not maintained open with cartilaginous rings as in an amniote trachea and remains closed between pulmonary breathing cycles. Therefore, only the volume of the buccal cavity and any residual air in the lungs contribute to respiratory dead space in a lungfish. Our calculations suggest dead space volume in L. paradoxa being between 4.1 and 12.5% of VT, values significantly lower than in R. schneideri. These low values may be due to the relatively small buccal cavity with a relatively constant volume when compared to the toad. Da Silva et al. (2017) estimated maximum buccal breath volume to be around 15 mL kg−1 in L. paradoxa, a value much smaller than the 20–50 mL kg−1 reported for Lithobates catesbeianus (Gans et al., 1969) or the estimate of 25 mL kg−1 for R. schneideri (Fernandes et al., 2005). Assuming a mean dead space volume of 10% for L. paradoxa and a buccal cavity volume of 15 mL kg−1, only a volume of 1.5 mL kg−1 would constitute the dead space in this species. This small dead space volume would only be returned towards the lungs during the first buccal inspiration, whereas the following buccal inspirations would each pump 100% fresh air into the lungs. Assuming that lungfish exchange about 90% of pulmonary air during each ventilatory cycle (Bishop and Foxon, 1968; Brainerd, 1994; Harder, 2001; da Silva et al., 2017), a one kilogram animal would possess a lung volume of 37 mL, from which 33.3 mL would be expired and accordingly 3.7 mL would remain with the lungs. From the exhaled 33.3 mL, 1.5 mL would be mixed with inspired air and returned to the lungs. Consequently, immediately after a ventilatory cycle the lungs would contain 5.2 mL of used air and 31.8 mL of fresh air. Assuming a PIO2 of 150 mmHg and a PLO2 of 30 mmHg after a 10 min breath-hold, PLO2 immediately after inspiration and full mixture of pulmonary air, would be about 133 mmHg. And indeed, Johansen and Lenfant (1967) report PLO2 to be immediately after a ventilatory cycle at up to 140 mmHg in L. paradoxa and McMahon (1970) reports PLO2 of above 110 mmHg in P. aethiopicus, suggesting a nearly complete renewal of air during the two-stroke buccal pump mechanism. Moreover, considering that the Bohr VD equation deals with dilution of gas volumes, our data suggest that the degree of gas mixing within the buccal cavity was limited. If the following assumptions are considered: (i) buccal cavity volume ~ 15 mL kg−1, (ii) the gas expelled from the lungs mixes with the buccal gas, (iii) During non-ventilatory periods and immediately before
beginning of or before an expiration (corresponding to an opening of the mouth at the beginning of the ventilatory cycle; Brainerd et al., 1993), as well as immediately after a first buccal inspiration, they were not seen during expiratory airflow. We recorded a constant negative pressure being maintained within the buccal cavity (Fig. 1), and the accompanying ventilatory recordings are similar to the constant outflow of air seen in a previous study (da Silva et al., 2017). The presence of a constant negative pressure simultaneously to expiratory airflow seems counterintuitive, but could be explained by the fact that L. paradoxa possesses two pairs of internal connected nares (nares perforatae; Hyrtl, 1843), that are clearly visible during a ventilatory cycle (Goeldi, 1898). The outgoing airflow could be guided by a jet stream from the lungs to the posterior pair of internal nares, assisted by a series of longitudinal folds along the dorsal part of the mouth (Fig. 3). The expiratory flow leaving the animal through the nares could therefore be separated from the inspiratory flow occurring through the mouth, and the negative pressure measured could be explained by the expansion of the buccal cavity. As shown by Brainerd (1994), buccal diameter increases in L. paradoxa concomitantly with a decrease in lung diameter, and even after the lung has reached its minimum diameter of 40%, the buccal cavity diameter continues to increase. Once lung diameter stops decreasing, buccal cavity still increases some 30% in diameter (Brainerd, 1994). These findings suggest that expiratory airflow must occur through an expanding buccal cavity, resulting in the mixing of fresh and exhaled air. Our recordings suggest that the constant increase in buccal volume with the accompanying outflow of air, maintains pressure in the buccal cavity constantly sub-atmospheric. At point ‘c’ in Fig. 1, buccal pressure starts to rise sharply, becoming positive, but expiratory airflow continues. This suggests that at the first compression of the buccal cavity, some air is being expelled through the mouth, whereas the remaining air is being pumped into the lungs. The following expansions and compressions of the buccal cavity result in buccal cavity pressure oscillating between increasingly positive and small negative pressures. The increase in buccal pressure throughout the sequence of lung inflations could be explained in that as the maximal volume is approached, the total compliance of the system decreases. The buccal pressure oscillations observed at the end of the ventilatory cycle may be associated to the expulsion of air from the buccal cavity through the opercular openings (G.S.F·S, personnel observation; Goeldi, 1898; Bishop and Foxon, 1968), once the lungs have been filled completely. Ventilation data obtained in the present study are comparable to results obtained in previous studies using L. paradoxa at 25 °C (Table 3) and further confirm that V̇ E is being adjusted by changing fR, since expiratory VT is typically maintained at 25.0 ± 4.6 mL BTPS kg−1. 163
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ventilation, the buccal PCO2 is nearly zero; then we would expect to find PBCO2 during a ventilatory cycle (end-tidal CO2) to be much lower than described in the present study (~ 19 mmHg). Rather, we find a very small lung-buccal PCO2 difference (Table 1), which supports the idea of a near complete replacement of lung air during one ventilatory cycle. These data are in agreement with Fernandes et al. (2005) for the toads. In conclusion, L. paradoxa is able to renew almost all of its pulmonary air during one ventilatory cycle, releasing most of the air to the outside during the expansion of the buccal cavity. The degree of mixing within the buccal cavity is very small, as evidenced by the small dead space calculated, suggesting the presence of a ‘jet stream’ of used air leaving the lungs and rapidly passing through a slowly expanding buccal cavity. A jet stream mechanism may have evolved independently in lungfish and some anuran amphibians. But if considered being a plesiomorphic breathing mechanism of buccal pump breathing sarcopterygians, jet streaming would have significant implications on the evolution of air breathing in early tetrapods.
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Conflict of interests The authors confirm that there is no conflict of interest Acknowledgements Funding: This study was financially supported by the Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq), through the National Institute of Science and Technology on Comparative Physiology (INCT em Fisiologia Comparada, proceeding nos. 2008/ 57712-4 and 573921/2008-3). G.S.F. da Silva was supported by a Young Investigator Award (FAPESP no 2013/17606-9 and 2014/ 12190-1) and is currently supported by CNPq (403769/2016-7). W. J. Minto was the recipient of a FAPESP postdoc fellowship (no. 2009/ 53916-7). References Amin-Naves, J., Giusti, H., Glass, M.L., 2004. Effects of acute temperature changes on aerial and aquatic gas exchange, pulmonary ventilation and blood gas status in the south American lungfish, Lepidosiren paradoxa. Comp. Biochem. Physiol A 138, 133–139. Amin-Naves, J., Giusti, H., Hoffmann, A., Glass, M.L., 2007a. Components to the acid–base related ventilatory drives in the south American lungfish Lepidosiren paradoxa. Respir. Physiol. Neurobiol. 155, 35–40. Amin-Naves, J., Giusti, H., Hoffmann, A., Glass, M.L., 2007b. Central ventilatory control in the south American lungfish, Lepidosiren paradoxa: distinct responses to pH and CO2. J. Comp. Physiol. B. 177, 529–534. Bassi, M., Klein, W., Fernandes, M.N., Perry, S.F., Glass, M.L., 2005. Pulmonary oxygen diffusing capacity of the south American lungfish Lepidosiren paradoxa: physiological values by the Bohr integration method. Physiol. Biochem. Zool. 78, 560–569. Bassi, M., Giusti, H., da Silva, G.S., Amin-Naves, J., Glass, M.L., 2010. Blood gases and cardiovascular shunt in the south American lungfish (Lepidosiren paradoxa) during normoxia and hyperoxia. Respir. Physiol. Neurobiol. 173, 47–50. Bishop, I.R., Foxon, G.E.H., 1968. The mechanism of breathing in the south American lungfish, Lepidosiren paradoxa; a radiological study. J. Zool. Lond. 154, 263–271. Brainerd, E.L., 1994. The evolution of lung gill bimodal breathing and the homology of vertebrate respiratory pumps. Amer. Zool. 34, 289–299. Brainerd, E.L., Ferry-Graham, L.A., 2006. Mechanics of respiratory pumps. In: Shadwick, R.E., Lauder, G.V. (Eds.), Fish Biomechanics Fish Physiology series. 23. pp. 1–28. Brainerd, E.L., Ditelberg, J.S., Brambçe, D.M., 1993. Lung ventilation in salamanders and the evolution of vertebrate air-breathing mechanisms. Biol. J. Linn. Soc. 49, 163–183. Brett, S.S., Shelton, G., 1979. Ventilatory mechanisms of the amphibian, Xernopus laevis; the role of the buccal force pump. J. Exp. Biol. 80, 251–269. Brinkmann, H., Venkatesh, B., Brenner, S., Meyer, A., 2004. Nuclear protein-coding genes support lungfish and not the coelacanth as the closest living relatives of land vertebrates. Proc. Natl. Acad. Sci. 101, 4900–4905. Clement, A.M., Long, J.A., 2010. Air-breathing adaptation in a marine Devonian lungfish. Biol. Lett. 6, 509–512. da Silva, G.S.F., Giusti, H., Sanchez, A.P., do Carmo, J.M., Glass, M.L., 2008. Aestivation in the south American lungfish, Lepidosiren paradoxa: effects on cardiovascular function, blood gases, osmolality and leptin levels. Respir. Physiol. Neurobiol. 164, 380–385. da Silva, G.S.F., Giusti, H., Branco, L.G.S., Glass, M.L., 2011. Combined ventilatory
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