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Distribution and mixing of a liquid bolus in pleural space
Respiratory Physiology & Neurobiology 150 (2006) 287–299
Distribution and mixing of a liquid bolus in pleural space Francesca Bodega, Claudio Tresoldi, Cristina Porta, Luciano Zocchi, Emilio Agostoni ∗ Istituto di Fisiologia Umana I, Universit`a di Milano, Via Mangiagalli 32, I-20133 Milano, Italy Accepted 27 April 2005
Abstract Distribution and mixing time of boluses with labeled albumin in pleural space of anesthetized, supine rabbits were determined by sampling pleural liquid at different times in various intercostal spaces (ics), and in cranial and caudal mediastinum. During sampling, lung and chest wall were kept apposed by lung inflation. This was not necessary in costo-phrenic sinus. Here, 10 min after injection, lung inflation increased concentration of labeled albumin by 50%. Lung inflation probably displaces some pleural liquid cranio-caudally, increasing labeled albumin concentration caudally to injection point (6th ics), and decreasing it cranially. Boluses of 0.1–1 ml did not preferentially reach mediastinal regions, as maintained by others. Time for an approximate mixing was ∼1 h for 0.1 ml, and ∼30 min for 1 ml. This relatively long mixing time does not substantially affect determination of contribution of lymphatic drainage through stomata to overall removal of labeled albumin from 0.3 ml hydrothoraces lasting 3 h [Bodega, F., Agostoni, E., 2004. Contribution of lymphatic drainage through stomata to albumin removal from pleural space. Respir. Physiol. Neurobiol. 142, 251–263]. © 2005 Elsevier B.V. All rights reserved. Keywords: Bolus distribution in pleural space; Labeled albumin, removal; Rabbits; Mixing time; Passive lung inflation; Pleural liquid; Displacement; Thickness
1. Introduction A small liquid bolus introduced into the pleural space seems to be quickly drawn to the nearest margins of the pulmonary lobes, where the lung recoil is greater than in the rest of the lung because of its local deformation caused by the tight fit between lung and chest wall under physiological conditions. This has been sug∗ Corresponding author. Tel.: +39 02 50315432; fax: +39 02 50315430. E-mail address: [email protected] (E. Agostoni).
1569-9048/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.resp.2005.04.022
gested by the following findings. When a small volume of saline solution is introduced into the pleural space of cats in the supine or prone posture, the thickness of the pleural liquid in the most dependent part of the space does not increase as it does when the volume injected is greater than 0.7 ml (∼0.25 ml/kg). This indirect information regarding the distribution of the bolus refers to the first few minutes because the thickness of the pleural liquid was measured on specimens obtained after quick-freezing of the chest a few minutes since the injection of the bolus (Agostoni and D’Angelo, 1969). The above hypothesis has been validated in
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dogs in the lateral posture with a different experimental approach (Zocchi et al., 1993). A small bolus of Ringer (0.5 ml, ∼0.03 ml/kg) with methylene blue was let into the pleural cavity through a cannula in the 10th intercostal space. Through “windows” of endothoracic fascia in the 8th and 6th intercostal space, the dye was observed to reach the caudal border of the lung within a few seconds, in some cases it also reached the margin between the middle and the caudal lobe. The distribution of the dye was assessed on autopsy 12–25 min after the entrance of the bolus; the visceral and parietal pleura were only stained in the parts corresponding to the caudal border of the lung, the margins between middle and caudal lobe, and the path between the entrance site and the dorsal region of the caudal border of the lung (Zocchi et al., 1993). In few other experiments (the study could not be completed because in practice dogs were no longer available for research), the bolus was let into the pleural space through the 6th intercostal space (three dogs in the lateral and one in the supine posture). The visceral and parietal pleura were only stained in the parts corresponding to the caudal border of the lung, the margins between middle and caudal lobe, and the paths between the entrance site and these regions (L. Zocchi, P.G. Agostoni and E. Agostoni, unpublished observations). This direct information regarding the distribution of a small bolus in dogs also refers to the first few minutes because thereafter the bolus has no further staining capability, as shown by the finding that the regions stained were the same 12 and 25 min after the entrance of the bolus. On the other hand, a different distribution of a small bolus during the first few minutes has been obtained by monitoring the movements of 99m Tcalbumin with a gamma camera interfaced with a computer. The small bolus (0.7–0.9 ml, ∼0.05–0.08 ml/kg) was slowly injected into the pleural cavity through the 2nd, 4th or 6th intercostal space of supine dogs. The gamma camera was placed horizontally over the chest, and thus provided a frontal projection of the activity of labeled albumin. According to the interpretation of the images given by the authors, labeled albumin preferentially reached the diaphragmatic and mediastinal lung regions within a few seconds, and then also reached the apical region (Miserocchi et al., 1984; Negrini et al., 1985). Moreover, the specific activity of the mediastinal and diaphragmatic
regions remained markedly greater than that of the other regions (including those close to the site of injection) for 2 h (Negrini et al., 1985). This result would indicate that a rough mixing of a small bolus is not achieved even after 2 h. This phenomenon would imply problems in determining the contribution of the lymphatic drainage through the stomata of the parietal pleura to the overall removal of labeled albumin from the pleural space because the stomata are quite unevenly distributed (Wang, 1975, 1985; Negrini et al., 1991). On the other hand, the images provided by the frontal projection of the activity of radiolabeled albumin may be quite deceptive because of the complex geometry of the pleural space and of its depth (see Section 4.1). The first aim of the present research was, therefore, to determine the distribution of a small bolus with labeled albumin in the pleural space of supine rabbits at various times, in order to establish whether or not it is similar to that obtained from the images provided by the frontal projection of the activity of radiolabeled albumin injected with a small bolus in supine dogs. To this end, we used an approach based on sampling pleural liquid in various sites, i.e., a completely different approach from the one described above. The second aim was to assess with the same experimental approach the time required for an approximate mixing of a small, medium, or large bolus in the pleural space of rabbits. This information is necessary to establish whether mixing time may be so long as to substantially interfere with the determination of the contribution made by the lymphatic drainage through the stomata to the overall removal of labeled albumin from small to medium hydrothoraces. This check will be made on data of a previous research in which this determination was made in 0.3 ml hydrothoraces lasting 3 h (Bodega and Agostoni, 2004).
2. Methods 2.1. Experimental protocol and series of experiments The experiments were performed on 58 supine rabbits (2.3–3.0 kg body weight, b.w.). The rabbits were purchased from “bmg allevamento”, Cividate al Piano (Bergamo) or from “G. Bettinardi”, Momo
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(Novara). Animal experimentation was authorized by the Ministry of Health by decree No. 60/03A issued according to Order of the Executive 116/92, in compliance with Directive 86/609/EC. The animals were anesthetized with a mixture of sodium pentobarbital (Sigma, 12 mg/ml) and urethane (Sigma, 150 mg/ml); the initial dose was 2 ml/kg intravenously, and small additional injections (0.2 ml/kg) were delivered when required to maintain an adequate level of anaesthesia. The trachea was cannulated, and connected to a heated Fleisch pneumotachygraph coupled to a Sanborn 270 differential pressure transducer to measure of air flow and tidal volume (obtained by electronic integration the flow signal by means of a HewlettPackard 8815A respiratory integrator). Air flow and tidal volume were recorded on a 7418 Hewlett-Packard thermopaper oscillograph throughout the experiment. Skin and muscles down to the intercostal muscles were removed from most of the right and left sides of the rib cage. On the right side, pleural “windows” were made at about mid-height in three of the following intercostal spaces (ics): 2nd, 5th, 7th, 8th, and 9th. All tissues, except the endhotoracic fascia, were removed over an area of ∼0.5 cm2 . A double thread loop was prepared in the intercostal muscles of the right 6th ics ∼0.5 cm below mid-height, and a stainless-steel cannula (1.4 mm o.d.; 0.9 mm i.d.) was inserted in the pleural space, and tightened to the intercostal muscles by one thread loop. The part of the cannula to be inserted had four holes, was slightly bent, and its closed, smooth tip pointed upwards. The cannula was connected to a glass syringe containing the solution to be injected. The injectate was Ringer solution (composition in mM: Na+ 139, K+ 5, Ca2+ 2.5, Mg2+ 1.5, Cl− 119, HCO3 − 29, d-glucose 5.6) containing 0.5 mg/ml bovine serum albumin conjugated with the fluorescent molecule Texas Red (Molecular Probes). Unlabeled albumin was added so that total albumin concentration in the solution was similar to that occurring in the pleural liquid under physiological conditions (10 mg/ml). A bolus of 0.1, 0.5 or 1 ml of the solution was slowly (∼0.1 ml in 5 s) injected into the pleural space; the cannula was then removed, and the second thread loop was tightened (Fig. 1). The 6th ics was chosen as injection point because it is far from lobar margins, is nearly midway along the cranio-caudal distance of the pleural space, and has been recently used in a research on the removal of labeled macromolecules from the pleural
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Fig. 1. Semischematic drawing of right lung lobes, rib cage, and part of diaphragm apposed to rib cage. Broken lines indicate lobe borders after passive lung inflation to ∼10 cm H2 O. Filled circle indicates injection point. Rib cage modified from Barone et al. (1973). Relationships between lung borders and intercostal spaces were checked in vivo.
space (Bodega and Agostoni, 2004; see Section 4.2). Breathing frequency and tidal volume were measured during the initial, middle, and final part of each experiment, and minute ventilation was computed. The three values were averaged to obtain the mean ventilation in a given experiment. The number of deep breaths was also measured. Ten, 30 or 50 min after the injection, the rabbit was killed by an intravenous injection of KCl. The lung was passively inflated to an alveolar pressure of ∼10 cm H2 O to keep the lung apposed to the chest wall during sampling of pleural liquid (see below). Alveolar pressure was measured by a pressure transducer (Hewlett-Packard 1280c) and recorded on the oscillograph. The transducer was connected to the trachea, and coupled to a Hewlett-Packard 8805 B carrier amplifier. Part of the left rib cage was then quickly removed to expose two small areas of the mediastinum. One ∼2 cm caudal to lung hilum, slightly below midheight, and the other one ∼0.5 cm cranial to lung hilum, above mid-height. In some experiments the lung was passively inflated after opening the left rib cage; in this case, the trachea was closed before opening the left rib cage. The area of the mediastinum cranial to the hilum can be reached only by displacing the pericardial sac with the surrounding fat slightly downwards and to the left. This operation requires some severing of tissues, and may, therefore, cause small holes in the mediastinal pleura and the entrance of air into the right pleural space, despite the passive inflation of the lung. This is probably caused by the traction exerted by the weight of the heart. When the volume of the air entered was more than a few little bubbles, the experiment was discarded.
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In order to establish whether the operation required to reach the cranial mediastinum affects the concentration of labeled albumin in the sample obtained in this area, we performed some experiments in which the cranial mediastinum was reached through the pericardial sac. In these experiments, the sternum was first cut along its transversal axis ∼1 cm cephalad of the xyphoid process. The ribs were then cut along their sternal ends so that the parietal pleura on both sides remained undamaged. Part of the sternum was removed by making a second transversal incision near the manubrium. The pericardium was then cut longitudinally, and the heart was displaced and supported. In this way, a small area of the right mediastinum craniad to lung hilum was visible through the parietal pericardium. The exposure of the mediastinal areas requires nearly 10 min with both approaches. In order to establish whether this post mortem delay affects the distribution of the labeled albumin concentration in the samples of pleural liquid (see Section 2.2), in some experiments sampling was performed first through the intercostal “windows”, and vice versa. Moreover, in some experiments only one mediastinal area was exposed so that the time was reduced. The concentration of labeled albumin in the various sites was not affected by the different delays. Finally, the concentration of labeled albumin in the cranial mediastinal region was similar with both approaches. At the end of the sampling procedure, the rabbit was placed in the left lateral posture 80◦ head up. The right pleural space was widely opened, and its liquid collected through a polyethylene tube connected to a 1 ml glass syringe. This liquid was collected for two purposes. First, to correct the concentration of labeled albumin in the microsamples of pleural liquid collected in the various sites (see Section 2.2). Second, to compare its labeled albumin concentration with the esti ). This virtual mated mixed, initial concentration (Cmix concentration is estimated from the quantity of labeled albumin injected and the initial volume of pleural liquid, which, in turn, is given by the volume of the bolus injected plus the physiological volume of the pleural liquid. The latter was computed from body weight as 0.30 × b.w.2/3 (Bodega and Agostoni, 2004). The above comparison is rough because when the posture of the rabbit is changed to collect pleural liquid, some liquid with a different concentration may be lost from the various openings made in the right pleural space
and the liquid collected, therefore, cannot be assumed to be well mixed. However, this comparison is useful to recognize rabbits with an abnormally large volume of pleural liquid. These rabbits, as well as those with evident pathological signs, were discarded. To study the effect of time and liquid volume on the distribution and mixing of the bolus injected, we performed five series of experiments: three series with an injection of 0.1 ml lasting 10, 30, and 50 min, one series with injection of 0.5 ml lasting 10 min, and one series with injection of 1 ml lasting 30 min. The sampling order was changed from one experiment to another. Several combinations of sampling sites were used, but no more than five sites were sampled in a given rabbit. Moreover, taking advantage of the fact that in supine rabbits the pleural pressure at mid-height of the zone of apposition between diaphragm and rib cage should not be subatmospheric (Perez et al., 1993), samples from this space were also taken before passive lung inflation in some rabbits in which a “window” was made in the 9th ics. Finally, for the reasons indicated under Section 3.1, in three extra rabbits in the supine posture we measured pleural liquid pressure in the 6th and 9th ics at the same height (about midway dorso-ventrally). The cannulas used were similar to the one used to inject the bolus in the above experiments. Each cannula was introduced under a layer of liquid to avoid the entry of air into the pleural space during the maneuver, and was connected to a Sanborn 268 differential pressure transducer through a polyethylene tubing and a three-way stopcock. The system was filled with Ringer solution. The transducer membrane was placed at the height of the holes of the cannula. The transducer was connected to a Hewlett-Packard 8805B carrier amplifier, and recordings were made as indicated above. After recording pleural liquid pressure under spontaneous breathing, the animal was paralyzed with pancuronium bromide (0.1 mg/kg, i.v.) and mechanically ventilated (Harvard Respirator 665) with a tidal volume and frequency similar to those occurring during spontaneous breathing. The animal was then disconnected from the respirator, and a passive lung inflation to about 10 cm H2 O was performed with a syringe. After another period of mechanical ventilation, passive lung inflation was repeated. The animal was then killed by intravenous injection of KCl, and after a few minutes passive lung inflation
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was performed twice. The pleural liquid pressure was recorded during all these experimental conditions.
were taken from the liquid collected in the right pleural space of four rabbits in which no bolus was injected.
2.2. Sampling procedure
2.3. Fluorescence measurements and statistics
In order to sample pleural liquid at various sites, a filter paper disk was inserted in the pleural space through a small incision made in the intercostal “windows”, and on the exposed areas of the mediastinum. The disk was left in the space for 15–20 s. Sampling in three intercostal sites required ∼4 min, and a similar time was required for sampling in the two mediastinal sites. The disks, 0.5 cm in diameter, were obtained from Whatman Quantitative Filter Paper, Grade 41. Before and immediately after sampling, each disk was weighed in the same test-tube, provided with a closing cap. The volume of liquid absorbed by the disk (3–6 l) was obtained by weight difference. An analytical balance (Gibertini, Crystal 100, Milan) with a resolution of 0.1 mg was used. Each tube (with disk) was weighed three times and the three values were averaged. The range of values for a given disk was never greater than 0.2 mg, i.e., <10% of the volume of liquid absorbed by the disks. After determining the volume of pleural liquid absorbed by the disk, 2.3 ml of Ringer were added in the tube. Then, the tube was sonicated for 10 min, incubated in an orbital shaker for 2 21 h, and then sonicated again for 10 min; thereafter, the fluorescence intensity of the solution was measured (see Section 2.3). In order to determine the fraction of labeled albumin absorbed by the disk that can be released into the solution at the end of the above incubation process, three disks for each experiment were soaked with 5 l of pleural liquid collected from the right pleural space at the end of the experiment (see Section 2.1). The disks were then incubated as described above, and the fluorescence intensity was measured. The three values were averaged, and the mean divided by the fluorescence intensity of the pleural liquid in which the disks were soaked. Preliminary experiments had shown that ∼94% of the labeled albumin taken up by the disk from the pleural liquid is released in the solution where the disk is incubated. This fraction was determined in each experiment, and used to correct the concentration of labeled albumin measured in the sample of a given disk in order to obtain the concentration of labeled albumin in the pleural liquid at the site where the disk was inserted. In order to determine the background fluorescence, disk samples
The fluorescence intensity in the liquid of incubation was measured by a spectrofluorophotometer (Shimadzu RF1501) (excitation 596 nm, emission 615 nm). A calibration curve was made for each experiment by diluting the injectate in Ringer solution. It was linear over the range 0–5 g/ml. A calibration factor was computed by linear regression through the points, and used to obtain the concentration of labeled albumin in the incubation liquid (Cinc ) of each disk. The concentration of labeled albumin in the liquid collected from each pleural site (C) was obtained from the corresponding Cinc , by taking into account the dilution undergone by the pleural liquid absorbed by the disk (Vabs ) in the incubation liquid (2300 l), and correcting this value for the fraction of labeled albumin absorbed by the disk that was released in the solution during incubation (Fr, see Section 2.2). Hence, C = (Cinc × 2300/Vabs )/Fr. Because the disks were often contaminated by a small amount of blood coming from the incision of the pleura, the volume of contaminating blood was estimated, and subtracted from the volume of liquid absorbed by the disk (see Section 2.2). After incubation, the red cells were hemolized (probably because of sonication), and the volume of the contaminating blood was then determined by reading the absorbance of hemoglobin in solution at 541 nm in a spectrophotometer (Zeiss PM2DL). A calibration curve was determined by adding 0.1–1.5 l of hemolized blood to 2.3 ml of Ringer. A calibration factor was obtained by linear regression through the points. The volume of contaminating blood was then calculated dividing the absorbance by the calibration factor. The sample was discarded if the volume of contaminating blood was greater than 20% of the volume sampled. The results are reported as mean values ± standard error (S.E.). The statistical difference among groups with passive lung inflation was assessed by the analysis of variance with post-hoc Bonferroni’s limitation for all the possible comparisons. Statistical significance was set at P < 0.05. When only two groups were to be compared (9th ics with and without passive lung inflation) the statistical difference between them was assessed with the unpaired t-test.
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3. Results 3.1. Bolus of 0.1 ml for 10, 30, and 50 min The concentration of labeled albumin in the various sampling sites of the pleural space 10, 30, and 50 min after the injection through the 6th intercostal space (ics) of 0.1 ml of Ringer-albumin with 500 g/ml of labeled albumin is reported in Table 1. The estimated mixed, initial concentration of labeled albumin in pleu ) is also reported in Table 1. This virtual ral liquid (Cmix concentration is computed from the quantity of labeled albumin injected and the initial volume of pleural liquid, which, in turn, is given by the volume of the bolus injected plus the physiological volume of pleural liquid estimated from body weight (see Section 2.1). Ten minutes after the injection, the labeled albumin concentration in sites far from the injection point (2nd ics, cranial and caudal mediastinum) was markedly and significantly smaller than that in sites close to injection point (5th, 7th, 8th ics), and in 9th ics, which is not so close. Moreover, the concentration in the 5th ics was significantly smaller than in the 7th, 8th, and 9th ics (Table 1). Hence, the concentration in the 5th ics was intermediate between that in the sites far from injection point and that in the 7th, 8th, and 9th ics, while the
concentration in the 9th ics was similar to that in the 7th and 8th ics (see below). The distribution of concentration does not seem to be affected by the number of deep breaths (one–three) occurring during the experiments, by ventilation or by sampling sequence (see Section 2.1). In four rabbits (not reported in Table 1) the labeled albumin concentration injected with the bolus was 1500 g/ml instead of 500 g/ml. In these rabbits, the distribution of the concentration of labeled albumin was similar to that reported in Table 1, while the concentration in a given site was approximately three times greater. The labeled albumin concentration in the 5th ics was significantly lower than that in the 7th ics (Table 1) despite the similar distance of these sites from the injection point (6th ics). Instead, the concentration in the 9th ics was similar to that in the 7th ics (Table 1) despite the greater distance of the former from the injection point. This striking finding may depend on the fact that the samples of pleural liquid were taken after passive lung inflation. Two phenomena should be considered in this connection. First, passive lung inflation tends to displace some pleural liquid caudally by shear flow because of the movement of the lung and of the diaphragm relative to the rib cage. In sites caudal to the injection point, this movement draws liquid with a
Table 1 Labeled albumin concentration (g/ml) in various sites of the pleural space 10, 30 or 50 min after injection through the 6th intercostal space of 0.1 ml of Ringer-albumin with 500 g/ml of labeled albumin Time (min)
n
10c
8d
30
8e
50c
8f
Body weight (kg)
2.68 ±0.05 2.63 ±0.07 2.56 ±0.06
Estimateda in mixed, initial pleural liquid
Sampling site Intercostal space 2nd cranial lobe
5th middle lobe
32.7 ±7.9 55.8 ±8.8 56.2 ±5.2
72.3 ±3.2 – – – –
Mediastinum 7th caudal lobe 112 ±5.7 81.8 ±6.4 65.8 ±3.4
8th caudal border 103 ±4.2 86.7 ±6.3 68.6 ±3.1
9th costophrenicb
9th sinus
Cranial Middle or cranial lobe
Caudal Accessory lobe
71.3 ±7.5 – – 55.4 ±7.1
108 ±10.0 – – 67.1 ±6.7
29.6 ±8.4 50.9 ±6.9 47.2 ±3.6
36.1 ±9.6 47.8 ±5.7 50.8 ±3.2
Values are mean ± S.E. a See text. b Before lung inflation. c Sampling sites were three–five in individual rabbit. d Except 9th ics before lung inflation in which n was 5. e Except caudal mediastinum in which n was 7. f Except 7th and 8th ics in which n was 10, and 9th ics before and after lung inflation in which n was 4.
74.0 ±0.8 74.6 ±1.1 75.7 ±1.0
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greater concentration of labeled albumin, and therefore, increases the previous concentration. Instead, in sites cranial to the injection point (except the apical region) it draws liquid with a smaller concentration of labeled albumin, and therefore, decreases the previous concentration. Second, passive lung inflation increases pleural pressure more in the lung zone than in the apposition zone as appears from data in rabbits by Perez et al. (1993). Because the data of Perez et al. were obtained in the lateral posture, and because the rib capsule was placed in the 11th or 12th rib (although the zone of apposition does not reach the 12th rib), we performed three experiments in which the pleural liquid pressure was measured in the 6th and 9th ics at the same height in the supine posture. While pleural liquid pressure, averaged over the breathing cycle, was 6–7 cm H2 O greater in apposition than in lung zone during spontaneous breathing, after passive lung inflation to ∼10 cm H2 O (either in the living paralyzed animal or some minutes post-mortem) pleural liquid pressure was essentially the same in both zones, namely about nil. Hence, the slow liquid flow from apposition to lung zone, which probably occurs during spontaneous breathing (Agostoni et al., 1989), no longer occurs after passive lung inflation. Therefore, the cranio-caudal shear flow, described by the first consideration, takes place unopposed. In order to test whether the above hypothesis is correct we took advantage of the fact that pleural pressure at mid-height of the zone of apposition in supine rabbits is not subatmospheric, and a sample of pleural liquid may, therefore, be taken without passive lung inflation. Hence, in five rabbits samples of pleural liquid in the 9th ics were also taken before passive lung inflation to ∼10 cm H2 O. The labeled albumin concentration of these five rabbits after lung inflation (107 ± 12.4 g/ml) was 50% greater (P < 0.05) than before lung inflation (71.3 ± 7.5 g/ml). This finding supports our hypothesis that passive lung inflation displaces some pleural liquid cranio-caudally. Because of this displacement of pleural liquid, the labeled albumin concentration measured after lung inflation is overestimated in sites caudal to the injection point, and underestimated in sites cranial to the injection point. As a consequence, because without passive lung inflation labeled albumin concentration in the 5th and 7th ics should be similar, the concentration in these, and in the 8th and 9th ics 10 min after the injection might roughly be 90, 90, 80, and 70 g/ml, respectively, instead of
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the values reported in Table 1. The concentration in the other sites should be affected to a minor extent by the phenomenon described above. Even taking this artifact into account, there is a marked difference in concentration between the sites close to and far from injection point. Thirty minutes after the injection the labeled albumin concentration in sites far from the injection point (2nd ics, cranial and caudal mediastinum) was still significantly smaller than in sites close to the injection point (7th and 8th ics). This difference, however, was less than half of the difference between the same sites 10 min after the injection (Table 1). Fifty minutes after the injection, the concentration of labeled albumin in sites far from the injection point was similar to that occurring after 30 min, while the concentration in sites close to the injection point was significantly smaller than that occurring after 30 min. Fifty minutes after the injection the concentration in the 2nd ics was no longer significantly smaller than in sites close to the injection point, while the concentration in the mediastinal sites was still significantly smaller than in sites close to the injection point. This difference, however, was ∼half of the difference between the same sites 30 min after the injection (Table 1). In order to determine the overestimate of the concentration caused by passive lung inflation (see above) after 50 min, we measured the concentration in the 9th ics of four rabbits before and after passive lung inflation. The concentration after lung inflation was 21% greater than before lung inflation, but this difference was not significant. Hence, without lung inflation the concentration in the 7th and 8th ics, 50 min after the injection might roughly be 60 g/ml, i.e., close to the concentration in sites far from the injection point (Table 1). Even after 50 min the distribution of concentration does not seem to be affected by the number of deep breaths (4–13) occurring during the experiments, ventilation or sampling sequence. The time course of the labeled albumin concentration in sites far from the injection point can be obtained from the data in Table 1, taking into account that the initial concentration is zero, and probably remains so during the first few minutes (Fig. 2, continuous line). The time course of the concentration in a site close to the injection point can also be drawn (Fig. 2, broken line). In this connection, it should be remembered that the labeled albumin concentration in the latter site
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Fig. 2. Time course of concentration of labeled albumin in pleural liquid of various sites after injection of a 0.1 ml bolus with tracer through 6th ics. Continuous line: concentration in sites far from injection point (2nd ics, cranial and caudal mediastinum); initial concentration was zero, and likely remained such during first few min. Broken line: concentration close to injection point (7th ics). Arrow indicates estimated mixed, initial concentration (see text).
is somewhat overestimated (see above). It, therefore, appears that with such a small bolus, mixing is essentially reached after ∼1 h (at least for the requirement of the second aim of this research, see Sections 1 and 4.2). 3.2. Bolus of 0.5 ml for 10 min and of 1 ml for 30 min The labeled albumin concentration in various sites of the pleural space 10 min after the injection through
the 6th ics of 0.5 ml of Ringer-albumin with 500 g/ml of labeled albumin is reported in Table 2. The concentration in sites far from injection point (2nd ics, cranial and caudal mediastinum) was significantly smaller than in sites close to the injection point (7th and 8th ics). The labeled albumin concentration in various sites of the pleural space 30 min after the injection through the 6th ics of 1 ml of Ringer-albumin with 500 g/ml of labeled albumin is also reported in Table 2. The concentration in sites far from injection point was not significantly different from that in sites close to the injection point. Hence, with a 1 ml bolus (0.38 ml/kg) mixing is essentially reached after 30 min, while with a 0.1 ml bolus it is reached after ∼1 h (see Section 3.1). Though the labeled albumin concentration was the same in the boluses of different volumes, the labeled albumin concentration in the pleural liquid is greater when the bolus injected is larger, because of the smaller dilution of the bolus by the physiological pleural liquid. Therefore, in order to compare the distribution of the labeled albumin concentration between boluses of different volume, the concentration must be expressed as a per cent of the corresponding Cmix (see Sections 2.1 and 3.1). Fig. 3 provides a synoptic view of the distribution of the labeled albumin concentration expressed . When analyzing this histogram, it should as % Cmix be remembered that the concentration in the 7th and 8th ics is somewhat overestimated because of the artifact produced by passive lung inflation, and that this phenomenon tends to vanish when the differences in concentration are small. It appears from this histogram that when the volume of the bolus is increased from 0.1 to 0.5 ml (10 min in both cases) the concentration distribution becomes more uniform, nearly as much as when
Table 2 Labeled albumin concentration (g/ml) in various sites of the pleural space 10 or 30 min after injection through the 6th intercostal space of 0.5 or 1 ml of Ringer-albumin, respectively, with 500 g/ml of labeled albumin Bolus volume (ml)
0.5 1.0
Time (min)
10 30
n
9b 5
Body weight (kg)
2.64 ± 0.11 2.65 ± 0.16
Estimateda in mixed, initial pleural liquid
Sampling site Intercostal space
Mediastinum
2nd Cranial lobe
7th Caudal lobe
8th Caudal border
Cranial Middle or cranial lobe
Caudal Accessory lobe
166 ± 16.9 302 ± 30.1
299 ± 13.4 310 ± 7.6
298 ± 12.2 331 ± 7.2
145 ± 26.4 308 ± 43.7
188 ± 31.4 290 ± 27.3
Values are mean ± S.E. a See text. b Except cranial mediastinum in which n was 7.
234 ± 3.1 318 ± 4.6
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Fig. 3. Concentration of labeled albumin in pleural liquid, expressed as per cent of estimated mixed, initial concentration (see Sections 2.1 and 3.1), at various times and with boluses of different volume. Concentration in 7th and 8th ics is somewhat overestimated (particularly at 10 min) because of passive lung inflation (see Section 3.1).
the time is increased from 10 to 30 min with the bolus of 0.1 ml. Moreover, when the volume of the bolus is increased from 0.1 to 1 ml (30 min in both cases), the concentration distribution becomes more uniform than when the 0.1 ml bolus is kept for 50 min.
4. Discussion 4.1. Bolus distribution The results show that in supine rabbits a small liquid bolus (0.04 ml/kg) injected into the pleural space through the 6th intercostal space (ics) does not preferentially reach the mediastinal regions, as it has been maintained for supine dogs on the basis of the images taken by a gamma camera placed horizontally over the chest, thus providing the frontal projection of the activity of 99m Tc-albumin injected with a small bolus (0.05–0.08 ml/kg) into the pleural space through the 6th ics (Miserocchi et al., 1984; Negrini et al., 1985). Indeed, in our experiments the labeled albumin concentration in mediastinal regions, 50 min after the injection, was not greater than in the regions close to the injection point, even taking into account the overestimation of the concentration in the 7th and 8th ics caused
by passive lung inflation (Table 1 and Fig. 2). This discrepancy might be due to a species difference, but this does not seem to be the case, because even during the first few minutes the distribution of a dye injected with a small bolus through the 6th ics space of dogs is different from that obtained with a frontal projection of the activity of radiolabeled albumin (see Section 1). It, therefore, seems likely that the discrepancy between the present data on rabbits and those obtained in dogs with radiolabeled albumin depends on the deceptive image provided by the frontal projection of the activity of the tracer. Indeed, the following points should be taken into account, as partially done by the authors (Miserocchi et al., 1984; Negrini et al., 1985). (1) For a given concentration of tracer per unit of pleural surface and a given distance from the scanning plane, the regional signal on the plane increases with the increase in the angle formed by that area of pleura with the scanning plane. (2) Radiation energy per unit surface of the scanning plane (and hence, signal intensity) decreases markedly with the distance between the plane and the source of radiation. Moreover, this attenuation is enhanced by the density of the tissue crossed by the radiation. (3) Signal intensity depends on the sum of the frontal projections of all the regions of pleura at various depths subtended by a given area of the scanning plane. In
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addition, the initial part of the lymphatics draining the pleural space are in the field scanned by the gamma camera. Therefore, the complex geometry of the pleural space in some regions combined with the above three points on radiation measurement defies an analysis of tracer concentration in various regions of the pleural space based on a projection of tracer activity. This conclusion is supported by the following consideration. According to the image provided by the frontal projection on the gamma camera, the specific activity of the tracer in the mediastinal region, and in the latero-diaphragmatic lung region (another region preferentially reached by the bolus according to the authors), 1 h after the injection is three–four times greater than in the other regions of the pleural space (including those surrounding the injection point). This situation persists throughout the next hour (see Fig. 4 in Negrini et al., 1985), during which there is only a slow decay of signal intensity in all regions (varying in rate among them), likely related to the removal of the tracer from the pleural space, and its dilution by the liquid filtering into the pleural space. According to the authors, these marked differences in labeled albumin concentration are due to the preferential movement of the bolus to regions where pleural liquid pressure at a given height is more subatmospheric (mediastinal and diaphragmatic regions). On the other hand, the persistence of the marked differences in signal from the various regions, despite breathing and cardiac movements, suggests that they do not depend on differences in tracer concentration in the various pleural regions, but mainly on the factors mentioned in the above three points on radiation measurement. Indeed, our results show that 50 min after the injection into the pleural space of a small bolus, the labeled albumin concentration only differs slightly among various regions of the pleural space (see Section 3.1). The study of the distribution in the pleural space of a small bolus with radiolabeled albumin (Miserocchi et al., 1984) was mainly intended to support the finding by the same authors that pleural liquid pressure in the mediastinal regions is more subatmospheric than in the costal region at the same height (Miserocchi et al., 1981, 1984). Indeed, this difference in pressure was questionable, because the lower pressure in the mediastinal region might be due to the distortion caused by the introduction of the cannula in this region (Agostoni and D’Angelo, 1991). The present finding
that a small bolus does not preferentially reach the mediastinal region casts further doubts on the more subatmospheric pressure in the mediastinal region. At this stage, it should be borne in mind that the topographical distribution of the bolus changes with the position of the point of its injection, and that our data only refer to injection through the 6th ics. For instance, if the bolus were injected into the 3rd ics, the labeled albumin concentration after 10 min should be greater in the 2nd ics and the cranial mediastinum than in the 7th, 8th ics, and the caudal mediastinum. Moreover, each approach has its own limitations. Our approach does not provide information on the labeled albumin concentration in the interlobar and diaphragmatic lung regions; this limitation, however, should not prevent the information required by the purposes of the present research from being obtained. Sampling in the mediastinal regions takes ∼14 min after the death of the animal, but this delay does not seem to appreciably affect the labeled albumin concentration (see Sections 2.1 and 3.1). Finally, our approach requires passive lung inflation for sampling (see Section 2.1), except for the zone of apposition between diaphragm and rib cage where, in rabbits, pleural pressure is not subatmospheric. Passive lung inflation seems to displace some pleural liquid cranio-caudally (see Section 3.1). This liquid displacement increases the concentration of labeled albumin in the zone of apposition. On the basis of this increase, an approximate correction of the concentration in the regions surrounding the injection point was made (see Section 3.1). Unfortunately, this effect of passive lung inflation was only measurable in one site. 4.2. Mixing of the bolus and removal of labeled albumin Complete mixing of a small bolus is never reached because after 30 min the quantity of labeled albumin removed from the pleural space becomes appreciable, and the overall concentration, therefore, decreases because of liquid filtration into the pleural space when the bolus is small (Bodega and Agostoni, 2004). As a consequence, the concentration in sites far from the injection point no longer increases. Another factor preventing a complete mixing is the uneven removal of labeled albumin produced by the lymphatic drainage through the stomata, which are unevenly distributed
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(Wang, 1975, 1985; Negrini et al., 1991). This way of removal alone does not change the labeled albumin concentration, but since some liquid is filtered into the pleural space, the labeled albumin concentration decreases. This decrease in concentration should become greater from the 2nd to the 7th and 8th ics, and from these to the mediastinal regions, because the stomata are lacking in the apical region, and are more concentrated in the mediastinal than in the middle intercostal region. This phenomenon does not appear before 30 min because the prevailing changes in concentration of labeled albumin during this period are those due to the mixing of the liquid of the regions close to the injection point with liquid of the other regions. After 30 min, these changes become smaller, and therefore, those due to the uneven distribution of the stomata become apparent. The contribution of the lymphatic drainage through the stomata of the parietal mesothelium to the overall removal of labeled albumin from the pleural space has recently been measured in anesthetized rabbits with a small (0.3 ml, 0.12 ml/kg) or large (6 ml, 2.4 ml/kg) hydrothorax of Ringer-albumin with labeled albumin and labeled dextran-2000 kDa (Bodega and Agostoni, 2004). This dextran was used as marker for liquid removal through the stomata because, owing to its size, it should essentially leave the pleural space only through the stomata. The quantity of labeled albumin removed by the lymphatic drainage through the stomata was determined from the product of the volume of liquid removed through them and the estimated mean concentration of labeled albumin in the pleural liquid during the experiment. The removal of labeled albumin through the stomata turned out to be 39% of the overall removal in the small hydrothoraces and 64% in the large ones. On the other hand, it could be argued that the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin in the small hydrothorax has been substantially underestimated because of the following considerations. (1) The distribution of stomata in the parietal pleura is quite uneven (see above), (2) the above experiments lasted 3 h, and (3) the mixing time of a small hydrothorax was unknown. The assessment in the present research of the time required for approximate mixing of small and medium boluses (injected through the 6th ics as in the previous research; Bodega and Agostoni, 2004) provided the information for an analysis of this prob-
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lem. The mixing times for boluses of 0.1, 0.5, and 1 ml are ∼1, 3/4, and 1/2 h, respectively (see Section 3.1). Hence, for the small hydrothorax of the previous research (0.3 ml) the mixing time might have been nearly 1 h, i.e., substantial compared with the duration of the experiment (3 h). Instead, for the large hydrothorax (6 ml) the mixing time might have been less than 10 min, i.e., almost negligible. Hence, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin might have been appreciably underestimated in the small, but not in the large hydrothorax. Nevertheless, taking into account that the 6th ics is about equidistant from the region without stomata (apical region) and those with the highest concentration of stomata (mediastinal and diaphragmatic ones) (Wang, 1975, 1985; Negrini et al., 1991), the above underestimation might be small. The removal of labeled albumin through the stomata in the small hydrothorax was 12.6 g in 3 h, which corresponds to 39% of the overall removal (31.9 g; Bodega and Agostoni, 2004). Because mixing time in this hydrothorax is somewhat less than 1 h (see above), the removal of labeled albumin through the stomata during the 2nd and 3rd hour should not be affected by this phenomenon, while removal during the 1st hour could be smaller because of the uneven distribution of labeled albumin combined with the uneven distribution of stomata. To be safe, in the following example, the removal of labeled albumin through the stomata during the 1st hour has been taken as ∼half of that occurring with uniform distribution of labeled albumin. Therefore, an approximate estimate of the removal of labeled albumin through the stomata during each of the 3 h may be 2.6 g for the 1st and 5 g for the 2nd and 3rd, i.e., 12.6 g for the whole period (for simplicity, we disregarded the small difference in removal among hours caused by the 15% decrease in labeled albumin concentration during the experiment; Bodega and Agostoni, 2004). Instead, in case of an instantaneous mixing, the removal of labeled albumin through the stomata during the 1st hour would be greater, being similar to the removal during the 2nd or 3rd hour, and therefore, the removal during the whole 3-h period would also be greater than that found experimentally (12.6 g). Hence, in this virtual case an estimate of the removal of labeled albumin through the stomata during each of the 3 h would be 5 g, i.e., 15 g for the whole period. That is, the uneven distribution of
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labeled albumin during the mixing time, combined with the uneven distribution of the stomata, should involve a smaller removal of labeled albumin through the stomata of 2.4 g. The loss of labeled albumin through the other mechanisms (transcytosis, convection and diffusion) is 19.3 g, being given by the overall removal (31.9 g) minus the removal through the stomata (12.6 g). Assuming that the loss of labeled albumin through the other mechanisms did not change because of the relatively long mixing time, the overall removal of labeled albumin in case of instantaneous mixing would be 15 + 19.3 = 34.3 g, instead of 31.9 g (see above). Hence, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin would be of 15 over 34.3 g, i.e., 44% instead of 39%. Therefore, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin from the small hydrothorax would have been underestimated by only 5%. Furthermore, if the removal of labeled albumin through the other mechanisms also decreased because of the relatively long mixing time of the small hydrothorax, with instantaneous mixing this removal would be greater, and therefore, the overall removal of labeled albumin would also be greater. As a consequence, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin would be less than 44%. That is, the underestimate would be less than 5%. The above analysis, therefore, supports the conclusion of the previous research (Bodega and Agostoni, 2004), namely that under physiological conditions (i.e., with a volume of liquid in the pleural space even smaller than that with the small hydrothorax) the lymphatic drainage through the stomata does not contribute most of protein and liquid removal from the pleural space, contrary to what has previously been maintained by others (for literature see Bodega and Agostoni, 2004). 4.3. Pleural liquid thickness The indirect evidence provided by the present research that passive lung inflation displaces some pleural liquid cranio-caudally suggests to reconsider the interpretation by Wang and Lai-Fook (1997) of their findings that pleural liquid thickness increases from the 2nd to the 8th ics in rabbits ventilated by positive pressure. To this end, one has first to recall a previous research by the same authors in which the thickness
of pleural liquid of rabbits ventilated with positive pressure was found to increase with the increase in ventilation (Wang and Lai-Fook, 1993). The pleural liquid thickness was measured by fluorescence videomicroscopy through a parietal pleura “window” in the 4th or 6th ics. According to their interpretation, the increase in pleural liquid thickness with the increase in ventilation is due to liquid displacement from the lobar margins, which would decrease power dissipation when the velocity of lung movements is increased (Wang and Lai-Fook, 1997). On the other hand, the above finding could simply be the consequence of positive pressure ventilation, which by increasing the pressure on the pleural surface displaces some pleural liquid beneath the “window”, which is more compliant than the rest of the chest wall (Agostoni and Zocchi, 1998). Indeed, during positive pressure ventilation the thickness of the pleural liquid beneath the “window” increased at each lung inflation (see Fig. 1 of Wang and Lai-Fook, 1993). In the following research on rabbits ventilated by positive pressure, in addition to the cranio-caudal increase in the pleural liquid thickness mentioned above, Wang and Lai-Fook (1997) found that a marked increase in ventilation did not increase the pleural liquid thickness in the 2nd ics, but did it moderately in the 5th ics, and markedly in the 8th ics. Because the relative velocity of the lung increases in the cranio-caudal direction for a given ventilation, they concluded that the increase in pleural liquid thickness from the cranial to the caudal regions has a functional relevance because it provides a uniform shear stress despite the different velocity of the lung in these regions. On the other hand, as pointed out by the authors, pleural liquid thickness is not a function of lung velocity alone, because with two values of ventilation the pleural liquid thickness was smaller in the 5th ics than in the 8th ics despite similar lung velocity. In connection with these findings and their interpretation the following considerations may be made. (1) The present research provides indirect evidence that passive lung inflation displaces some pleural liquid cranio-caudally. Therefore, during positive pressure breathing, pleural liquid thickness in the caudal ics should increase at each lung inflation. (2) The above-mentioned displacement of pleural liquid under the “window” caused by positive pressure breathing might be greater in the caudal than in the cranial “window” because the caudal ics are likely more
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compliant than the cranial ones because of the greater distance between the ribs. This may further contribute to the increase in pleural liquid thickness in the caudal “window” occurring during positive pressure breathing. (3) The increase in pleural liquid thickness with the increase in lung velocity would have a functional relevance if pleura lubrication were hydrodynamic, but a recent research has shown that it is boundary, i.e., frictional force is independent of velocity (D’Angelo et al., 2004). Therefore, an increase in thickness of the pleural liquid with the increase in lung velocity would not be of functional relevance.
Acknowledgment We thank R. Galli for skillful technical assistance.
References Agostoni, E., Agostoni, P.G., Zocchi, L., 1989. Pleural liquid pressure in the zone of apposition and in the lung zone. Respir. Physiol. 75, 357–370. Agostoni, E., D’Angelo, E., 1969. Thickness and pressure of the pleural liquid at various heights and with various hydrothoraces. Respir. Physiol. 6, 330–342. Agostoni, E., D’Angelo, E., 1991. Pleural liquid pressure. J. Appl. Physiol. 87, 393–403. Agostoni, E., Zocchi, L., 1998. Mechanical coupling and liquid exchanges in the pleural space. In: Antony, V.B. (Ed.), Clinics in Chest Medicine: Diseases of the Pleura. Saunders, Philadelphia, pp. 241–260. Barone, R., Pavaux, C., Blin, P.C., Cuq, P., 1973. Atlas d’anatomie du lapin. Masson & Cie , Paris, p. 18.
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Bodega, F., Agostoni, E., 2004. Contribution of lymphatic drainage through stomata to albumin removal from pleural space. Respir. Physiol. Neurobiol. 142, 251–263. D’Angelo, E., Loring, S.H., Gioia, M.E., Pecchiari, M., Moscheni, C., 2004. Friction and lubrication of pleural tissues. Respir. Physiol. Neurobiol. 142, 55–68. Miserocchi, G., Nakamura, T., Mariani, E., Negrini, D., 1981. Pleural liquid pressure over the interlobar mediastinal and diaphragmatic surfaces of the lung. Respir. Physiol. 46, 61–69. Miserocchi, G., Pistolesi, M., Miniati, M., Bellina, C.R., Negrini, D., Giuntini, C., 1984. Pleural liquid pressure gradients and intrapleural distribution of injected bolus. J. Appl. Physiol. 56, 526–532. Negrini, D., Pistolesi, M., Miniati, M., Bellina, R., Giuntini, C., Miserocchi, G., 1985. Regional protein absorption rates from the pleural cavity in dogs. J. Appl. Physiol. 58, 2062– 2067. Negrini, D., Mukenge, S., Del Fabbro, M., Gonano, C., Miserocchi, G., 1991. Distribution of diaphragmatic lymphatic stomata. J. Appl. Physiol. 70, 1544–1549. Perez, F., Fernandez, P., Hernaiz, M.I., Jackson, E.G., Lai-Fook, S.J., Boynton, B.R., 1993. Pleural pressure measured in the zone of apposition of diaphragm to rib cage in rabbits. Lung 171, 345–353. Wang, N.S., 1975. The preformed stomas connecting the pleural cavity and the lymphatics in the parietal pleura. Am. Rev. Respir. Dis. 111, 12–20. Wang, N.S., 1985. Mesothelial cells in situ. In: Chretien, J., Bignon, J., Hirsch, A. (Eds.), The Pleura in Health and Disease. Marcel Dekker Inc., New York, pp. 23–42. Wang, P.M., Lai-Fook, S.J., 1993. Effect of ventilation frequency and tidal volume on pleural space thickness in rabbits. J. Appl. Physiol. 75, 1836–1841. Wang, P.M., Lai-Fook, S.J., 1997. Effect of mechanical ventilation on regional variation of pleural liquid thickness in rabbits. Lung 175, 165–173. Zocchi, L., Agostoni, P.G., Agostoni, E., 1993. Intrapleural distribution of a liquid bolus let in the costophrenic sinus. Rend. Fis. Acc. Lincei Series 9, vol. 4, pp. 51–58.
Distribution and mixing of a liquid bolus in pleural space Francesca Bodega, Claudio Tresoldi, Cristina Porta, Luciano Zocchi, Emilio Agostoni ∗ Istituto di Fisiologia Umana I, Universit`a di Milano, Via Mangiagalli 32, I-20133 Milano, Italy Accepted 27 April 2005
Abstract Distribution and mixing time of boluses with labeled albumin in pleural space of anesthetized, supine rabbits were determined by sampling pleural liquid at different times in various intercostal spaces (ics), and in cranial and caudal mediastinum. During sampling, lung and chest wall were kept apposed by lung inflation. This was not necessary in costo-phrenic sinus. Here, 10 min after injection, lung inflation increased concentration of labeled albumin by 50%. Lung inflation probably displaces some pleural liquid cranio-caudally, increasing labeled albumin concentration caudally to injection point (6th ics), and decreasing it cranially. Boluses of 0.1–1 ml did not preferentially reach mediastinal regions, as maintained by others. Time for an approximate mixing was ∼1 h for 0.1 ml, and ∼30 min for 1 ml. This relatively long mixing time does not substantially affect determination of contribution of lymphatic drainage through stomata to overall removal of labeled albumin from 0.3 ml hydrothoraces lasting 3 h [Bodega, F., Agostoni, E., 2004. Contribution of lymphatic drainage through stomata to albumin removal from pleural space. Respir. Physiol. Neurobiol. 142, 251–263]. © 2005 Elsevier B.V. All rights reserved. Keywords: Bolus distribution in pleural space; Labeled albumin, removal; Rabbits; Mixing time; Passive lung inflation; Pleural liquid; Displacement; Thickness
1. Introduction A small liquid bolus introduced into the pleural space seems to be quickly drawn to the nearest margins of the pulmonary lobes, where the lung recoil is greater than in the rest of the lung because of its local deformation caused by the tight fit between lung and chest wall under physiological conditions. This has been sug∗ Corresponding author. Tel.: +39 02 50315432; fax: +39 02 50315430. E-mail address: [email protected] (E. Agostoni).
1569-9048/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.resp.2005.04.022
gested by the following findings. When a small volume of saline solution is introduced into the pleural space of cats in the supine or prone posture, the thickness of the pleural liquid in the most dependent part of the space does not increase as it does when the volume injected is greater than 0.7 ml (∼0.25 ml/kg). This indirect information regarding the distribution of the bolus refers to the first few minutes because the thickness of the pleural liquid was measured on specimens obtained after quick-freezing of the chest a few minutes since the injection of the bolus (Agostoni and D’Angelo, 1969). The above hypothesis has been validated in
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dogs in the lateral posture with a different experimental approach (Zocchi et al., 1993). A small bolus of Ringer (0.5 ml, ∼0.03 ml/kg) with methylene blue was let into the pleural cavity through a cannula in the 10th intercostal space. Through “windows” of endothoracic fascia in the 8th and 6th intercostal space, the dye was observed to reach the caudal border of the lung within a few seconds, in some cases it also reached the margin between the middle and the caudal lobe. The distribution of the dye was assessed on autopsy 12–25 min after the entrance of the bolus; the visceral and parietal pleura were only stained in the parts corresponding to the caudal border of the lung, the margins between middle and caudal lobe, and the path between the entrance site and the dorsal region of the caudal border of the lung (Zocchi et al., 1993). In few other experiments (the study could not be completed because in practice dogs were no longer available for research), the bolus was let into the pleural space through the 6th intercostal space (three dogs in the lateral and one in the supine posture). The visceral and parietal pleura were only stained in the parts corresponding to the caudal border of the lung, the margins between middle and caudal lobe, and the paths between the entrance site and these regions (L. Zocchi, P.G. Agostoni and E. Agostoni, unpublished observations). This direct information regarding the distribution of a small bolus in dogs also refers to the first few minutes because thereafter the bolus has no further staining capability, as shown by the finding that the regions stained were the same 12 and 25 min after the entrance of the bolus. On the other hand, a different distribution of a small bolus during the first few minutes has been obtained by monitoring the movements of 99m Tcalbumin with a gamma camera interfaced with a computer. The small bolus (0.7–0.9 ml, ∼0.05–0.08 ml/kg) was slowly injected into the pleural cavity through the 2nd, 4th or 6th intercostal space of supine dogs. The gamma camera was placed horizontally over the chest, and thus provided a frontal projection of the activity of labeled albumin. According to the interpretation of the images given by the authors, labeled albumin preferentially reached the diaphragmatic and mediastinal lung regions within a few seconds, and then also reached the apical region (Miserocchi et al., 1984; Negrini et al., 1985). Moreover, the specific activity of the mediastinal and diaphragmatic
regions remained markedly greater than that of the other regions (including those close to the site of injection) for 2 h (Negrini et al., 1985). This result would indicate that a rough mixing of a small bolus is not achieved even after 2 h. This phenomenon would imply problems in determining the contribution of the lymphatic drainage through the stomata of the parietal pleura to the overall removal of labeled albumin from the pleural space because the stomata are quite unevenly distributed (Wang, 1975, 1985; Negrini et al., 1991). On the other hand, the images provided by the frontal projection of the activity of radiolabeled albumin may be quite deceptive because of the complex geometry of the pleural space and of its depth (see Section 4.1). The first aim of the present research was, therefore, to determine the distribution of a small bolus with labeled albumin in the pleural space of supine rabbits at various times, in order to establish whether or not it is similar to that obtained from the images provided by the frontal projection of the activity of radiolabeled albumin injected with a small bolus in supine dogs. To this end, we used an approach based on sampling pleural liquid in various sites, i.e., a completely different approach from the one described above. The second aim was to assess with the same experimental approach the time required for an approximate mixing of a small, medium, or large bolus in the pleural space of rabbits. This information is necessary to establish whether mixing time may be so long as to substantially interfere with the determination of the contribution made by the lymphatic drainage through the stomata to the overall removal of labeled albumin from small to medium hydrothoraces. This check will be made on data of a previous research in which this determination was made in 0.3 ml hydrothoraces lasting 3 h (Bodega and Agostoni, 2004).
2. Methods 2.1. Experimental protocol and series of experiments The experiments were performed on 58 supine rabbits (2.3–3.0 kg body weight, b.w.). The rabbits were purchased from “bmg allevamento”, Cividate al Piano (Bergamo) or from “G. Bettinardi”, Momo
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(Novara). Animal experimentation was authorized by the Ministry of Health by decree No. 60/03A issued according to Order of the Executive 116/92, in compliance with Directive 86/609/EC. The animals were anesthetized with a mixture of sodium pentobarbital (Sigma, 12 mg/ml) and urethane (Sigma, 150 mg/ml); the initial dose was 2 ml/kg intravenously, and small additional injections (0.2 ml/kg) were delivered when required to maintain an adequate level of anaesthesia. The trachea was cannulated, and connected to a heated Fleisch pneumotachygraph coupled to a Sanborn 270 differential pressure transducer to measure of air flow and tidal volume (obtained by electronic integration the flow signal by means of a HewlettPackard 8815A respiratory integrator). Air flow and tidal volume were recorded on a 7418 Hewlett-Packard thermopaper oscillograph throughout the experiment. Skin and muscles down to the intercostal muscles were removed from most of the right and left sides of the rib cage. On the right side, pleural “windows” were made at about mid-height in three of the following intercostal spaces (ics): 2nd, 5th, 7th, 8th, and 9th. All tissues, except the endhotoracic fascia, were removed over an area of ∼0.5 cm2 . A double thread loop was prepared in the intercostal muscles of the right 6th ics ∼0.5 cm below mid-height, and a stainless-steel cannula (1.4 mm o.d.; 0.9 mm i.d.) was inserted in the pleural space, and tightened to the intercostal muscles by one thread loop. The part of the cannula to be inserted had four holes, was slightly bent, and its closed, smooth tip pointed upwards. The cannula was connected to a glass syringe containing the solution to be injected. The injectate was Ringer solution (composition in mM: Na+ 139, K+ 5, Ca2+ 2.5, Mg2+ 1.5, Cl− 119, HCO3 − 29, d-glucose 5.6) containing 0.5 mg/ml bovine serum albumin conjugated with the fluorescent molecule Texas Red (Molecular Probes). Unlabeled albumin was added so that total albumin concentration in the solution was similar to that occurring in the pleural liquid under physiological conditions (10 mg/ml). A bolus of 0.1, 0.5 or 1 ml of the solution was slowly (∼0.1 ml in 5 s) injected into the pleural space; the cannula was then removed, and the second thread loop was tightened (Fig. 1). The 6th ics was chosen as injection point because it is far from lobar margins, is nearly midway along the cranio-caudal distance of the pleural space, and has been recently used in a research on the removal of labeled macromolecules from the pleural
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Fig. 1. Semischematic drawing of right lung lobes, rib cage, and part of diaphragm apposed to rib cage. Broken lines indicate lobe borders after passive lung inflation to ∼10 cm H2 O. Filled circle indicates injection point. Rib cage modified from Barone et al. (1973). Relationships between lung borders and intercostal spaces were checked in vivo.
space (Bodega and Agostoni, 2004; see Section 4.2). Breathing frequency and tidal volume were measured during the initial, middle, and final part of each experiment, and minute ventilation was computed. The three values were averaged to obtain the mean ventilation in a given experiment. The number of deep breaths was also measured. Ten, 30 or 50 min after the injection, the rabbit was killed by an intravenous injection of KCl. The lung was passively inflated to an alveolar pressure of ∼10 cm H2 O to keep the lung apposed to the chest wall during sampling of pleural liquid (see below). Alveolar pressure was measured by a pressure transducer (Hewlett-Packard 1280c) and recorded on the oscillograph. The transducer was connected to the trachea, and coupled to a Hewlett-Packard 8805 B carrier amplifier. Part of the left rib cage was then quickly removed to expose two small areas of the mediastinum. One ∼2 cm caudal to lung hilum, slightly below midheight, and the other one ∼0.5 cm cranial to lung hilum, above mid-height. In some experiments the lung was passively inflated after opening the left rib cage; in this case, the trachea was closed before opening the left rib cage. The area of the mediastinum cranial to the hilum can be reached only by displacing the pericardial sac with the surrounding fat slightly downwards and to the left. This operation requires some severing of tissues, and may, therefore, cause small holes in the mediastinal pleura and the entrance of air into the right pleural space, despite the passive inflation of the lung. This is probably caused by the traction exerted by the weight of the heart. When the volume of the air entered was more than a few little bubbles, the experiment was discarded.
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In order to establish whether the operation required to reach the cranial mediastinum affects the concentration of labeled albumin in the sample obtained in this area, we performed some experiments in which the cranial mediastinum was reached through the pericardial sac. In these experiments, the sternum was first cut along its transversal axis ∼1 cm cephalad of the xyphoid process. The ribs were then cut along their sternal ends so that the parietal pleura on both sides remained undamaged. Part of the sternum was removed by making a second transversal incision near the manubrium. The pericardium was then cut longitudinally, and the heart was displaced and supported. In this way, a small area of the right mediastinum craniad to lung hilum was visible through the parietal pericardium. The exposure of the mediastinal areas requires nearly 10 min with both approaches. In order to establish whether this post mortem delay affects the distribution of the labeled albumin concentration in the samples of pleural liquid (see Section 2.2), in some experiments sampling was performed first through the intercostal “windows”, and vice versa. Moreover, in some experiments only one mediastinal area was exposed so that the time was reduced. The concentration of labeled albumin in the various sites was not affected by the different delays. Finally, the concentration of labeled albumin in the cranial mediastinal region was similar with both approaches. At the end of the sampling procedure, the rabbit was placed in the left lateral posture 80◦ head up. The right pleural space was widely opened, and its liquid collected through a polyethylene tube connected to a 1 ml glass syringe. This liquid was collected for two purposes. First, to correct the concentration of labeled albumin in the microsamples of pleural liquid collected in the various sites (see Section 2.2). Second, to compare its labeled albumin concentration with the esti ). This virtual mated mixed, initial concentration (Cmix concentration is estimated from the quantity of labeled albumin injected and the initial volume of pleural liquid, which, in turn, is given by the volume of the bolus injected plus the physiological volume of the pleural liquid. The latter was computed from body weight as 0.30 × b.w.2/3 (Bodega and Agostoni, 2004). The above comparison is rough because when the posture of the rabbit is changed to collect pleural liquid, some liquid with a different concentration may be lost from the various openings made in the right pleural space
and the liquid collected, therefore, cannot be assumed to be well mixed. However, this comparison is useful to recognize rabbits with an abnormally large volume of pleural liquid. These rabbits, as well as those with evident pathological signs, were discarded. To study the effect of time and liquid volume on the distribution and mixing of the bolus injected, we performed five series of experiments: three series with an injection of 0.1 ml lasting 10, 30, and 50 min, one series with injection of 0.5 ml lasting 10 min, and one series with injection of 1 ml lasting 30 min. The sampling order was changed from one experiment to another. Several combinations of sampling sites were used, but no more than five sites were sampled in a given rabbit. Moreover, taking advantage of the fact that in supine rabbits the pleural pressure at mid-height of the zone of apposition between diaphragm and rib cage should not be subatmospheric (Perez et al., 1993), samples from this space were also taken before passive lung inflation in some rabbits in which a “window” was made in the 9th ics. Finally, for the reasons indicated under Section 3.1, in three extra rabbits in the supine posture we measured pleural liquid pressure in the 6th and 9th ics at the same height (about midway dorso-ventrally). The cannulas used were similar to the one used to inject the bolus in the above experiments. Each cannula was introduced under a layer of liquid to avoid the entry of air into the pleural space during the maneuver, and was connected to a Sanborn 268 differential pressure transducer through a polyethylene tubing and a three-way stopcock. The system was filled with Ringer solution. The transducer membrane was placed at the height of the holes of the cannula. The transducer was connected to a Hewlett-Packard 8805B carrier amplifier, and recordings were made as indicated above. After recording pleural liquid pressure under spontaneous breathing, the animal was paralyzed with pancuronium bromide (0.1 mg/kg, i.v.) and mechanically ventilated (Harvard Respirator 665) with a tidal volume and frequency similar to those occurring during spontaneous breathing. The animal was then disconnected from the respirator, and a passive lung inflation to about 10 cm H2 O was performed with a syringe. After another period of mechanical ventilation, passive lung inflation was repeated. The animal was then killed by intravenous injection of KCl, and after a few minutes passive lung inflation
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was performed twice. The pleural liquid pressure was recorded during all these experimental conditions.
were taken from the liquid collected in the right pleural space of four rabbits in which no bolus was injected.
2.2. Sampling procedure
2.3. Fluorescence measurements and statistics
In order to sample pleural liquid at various sites, a filter paper disk was inserted in the pleural space through a small incision made in the intercostal “windows”, and on the exposed areas of the mediastinum. The disk was left in the space for 15–20 s. Sampling in three intercostal sites required ∼4 min, and a similar time was required for sampling in the two mediastinal sites. The disks, 0.5 cm in diameter, were obtained from Whatman Quantitative Filter Paper, Grade 41. Before and immediately after sampling, each disk was weighed in the same test-tube, provided with a closing cap. The volume of liquid absorbed by the disk (3–6 l) was obtained by weight difference. An analytical balance (Gibertini, Crystal 100, Milan) with a resolution of 0.1 mg was used. Each tube (with disk) was weighed three times and the three values were averaged. The range of values for a given disk was never greater than 0.2 mg, i.e., <10% of the volume of liquid absorbed by the disks. After determining the volume of pleural liquid absorbed by the disk, 2.3 ml of Ringer were added in the tube. Then, the tube was sonicated for 10 min, incubated in an orbital shaker for 2 21 h, and then sonicated again for 10 min; thereafter, the fluorescence intensity of the solution was measured (see Section 2.3). In order to determine the fraction of labeled albumin absorbed by the disk that can be released into the solution at the end of the above incubation process, three disks for each experiment were soaked with 5 l of pleural liquid collected from the right pleural space at the end of the experiment (see Section 2.1). The disks were then incubated as described above, and the fluorescence intensity was measured. The three values were averaged, and the mean divided by the fluorescence intensity of the pleural liquid in which the disks were soaked. Preliminary experiments had shown that ∼94% of the labeled albumin taken up by the disk from the pleural liquid is released in the solution where the disk is incubated. This fraction was determined in each experiment, and used to correct the concentration of labeled albumin measured in the sample of a given disk in order to obtain the concentration of labeled albumin in the pleural liquid at the site where the disk was inserted. In order to determine the background fluorescence, disk samples
The fluorescence intensity in the liquid of incubation was measured by a spectrofluorophotometer (Shimadzu RF1501) (excitation 596 nm, emission 615 nm). A calibration curve was made for each experiment by diluting the injectate in Ringer solution. It was linear over the range 0–5 g/ml. A calibration factor was computed by linear regression through the points, and used to obtain the concentration of labeled albumin in the incubation liquid (Cinc ) of each disk. The concentration of labeled albumin in the liquid collected from each pleural site (C) was obtained from the corresponding Cinc , by taking into account the dilution undergone by the pleural liquid absorbed by the disk (Vabs ) in the incubation liquid (2300 l), and correcting this value for the fraction of labeled albumin absorbed by the disk that was released in the solution during incubation (Fr, see Section 2.2). Hence, C = (Cinc × 2300/Vabs )/Fr. Because the disks were often contaminated by a small amount of blood coming from the incision of the pleura, the volume of contaminating blood was estimated, and subtracted from the volume of liquid absorbed by the disk (see Section 2.2). After incubation, the red cells were hemolized (probably because of sonication), and the volume of the contaminating blood was then determined by reading the absorbance of hemoglobin in solution at 541 nm in a spectrophotometer (Zeiss PM2DL). A calibration curve was determined by adding 0.1–1.5 l of hemolized blood to 2.3 ml of Ringer. A calibration factor was obtained by linear regression through the points. The volume of contaminating blood was then calculated dividing the absorbance by the calibration factor. The sample was discarded if the volume of contaminating blood was greater than 20% of the volume sampled. The results are reported as mean values ± standard error (S.E.). The statistical difference among groups with passive lung inflation was assessed by the analysis of variance with post-hoc Bonferroni’s limitation for all the possible comparisons. Statistical significance was set at P < 0.05. When only two groups were to be compared (9th ics with and without passive lung inflation) the statistical difference between them was assessed with the unpaired t-test.
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3. Results 3.1. Bolus of 0.1 ml for 10, 30, and 50 min The concentration of labeled albumin in the various sampling sites of the pleural space 10, 30, and 50 min after the injection through the 6th intercostal space (ics) of 0.1 ml of Ringer-albumin with 500 g/ml of labeled albumin is reported in Table 1. The estimated mixed, initial concentration of labeled albumin in pleu ) is also reported in Table 1. This virtual ral liquid (Cmix concentration is computed from the quantity of labeled albumin injected and the initial volume of pleural liquid, which, in turn, is given by the volume of the bolus injected plus the physiological volume of pleural liquid estimated from body weight (see Section 2.1). Ten minutes after the injection, the labeled albumin concentration in sites far from the injection point (2nd ics, cranial and caudal mediastinum) was markedly and significantly smaller than that in sites close to injection point (5th, 7th, 8th ics), and in 9th ics, which is not so close. Moreover, the concentration in the 5th ics was significantly smaller than in the 7th, 8th, and 9th ics (Table 1). Hence, the concentration in the 5th ics was intermediate between that in the sites far from injection point and that in the 7th, 8th, and 9th ics, while the
concentration in the 9th ics was similar to that in the 7th and 8th ics (see below). The distribution of concentration does not seem to be affected by the number of deep breaths (one–three) occurring during the experiments, by ventilation or by sampling sequence (see Section 2.1). In four rabbits (not reported in Table 1) the labeled albumin concentration injected with the bolus was 1500 g/ml instead of 500 g/ml. In these rabbits, the distribution of the concentration of labeled albumin was similar to that reported in Table 1, while the concentration in a given site was approximately three times greater. The labeled albumin concentration in the 5th ics was significantly lower than that in the 7th ics (Table 1) despite the similar distance of these sites from the injection point (6th ics). Instead, the concentration in the 9th ics was similar to that in the 7th ics (Table 1) despite the greater distance of the former from the injection point. This striking finding may depend on the fact that the samples of pleural liquid were taken after passive lung inflation. Two phenomena should be considered in this connection. First, passive lung inflation tends to displace some pleural liquid caudally by shear flow because of the movement of the lung and of the diaphragm relative to the rib cage. In sites caudal to the injection point, this movement draws liquid with a
Table 1 Labeled albumin concentration (g/ml) in various sites of the pleural space 10, 30 or 50 min after injection through the 6th intercostal space of 0.1 ml of Ringer-albumin with 500 g/ml of labeled albumin Time (min)
n
10c
8d
30
8e
50c
8f
Body weight (kg)
2.68 ±0.05 2.63 ±0.07 2.56 ±0.06
Estimateda in mixed, initial pleural liquid
Sampling site Intercostal space 2nd cranial lobe
5th middle lobe
32.7 ±7.9 55.8 ±8.8 56.2 ±5.2
72.3 ±3.2 – – – –
Mediastinum 7th caudal lobe 112 ±5.7 81.8 ±6.4 65.8 ±3.4
8th caudal border 103 ±4.2 86.7 ±6.3 68.6 ±3.1
9th costophrenicb
9th sinus
Cranial Middle or cranial lobe
Caudal Accessory lobe
71.3 ±7.5 – – 55.4 ±7.1
108 ±10.0 – – 67.1 ±6.7
29.6 ±8.4 50.9 ±6.9 47.2 ±3.6
36.1 ±9.6 47.8 ±5.7 50.8 ±3.2
Values are mean ± S.E. a See text. b Before lung inflation. c Sampling sites were three–five in individual rabbit. d Except 9th ics before lung inflation in which n was 5. e Except caudal mediastinum in which n was 7. f Except 7th and 8th ics in which n was 10, and 9th ics before and after lung inflation in which n was 4.
74.0 ±0.8 74.6 ±1.1 75.7 ±1.0
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greater concentration of labeled albumin, and therefore, increases the previous concentration. Instead, in sites cranial to the injection point (except the apical region) it draws liquid with a smaller concentration of labeled albumin, and therefore, decreases the previous concentration. Second, passive lung inflation increases pleural pressure more in the lung zone than in the apposition zone as appears from data in rabbits by Perez et al. (1993). Because the data of Perez et al. were obtained in the lateral posture, and because the rib capsule was placed in the 11th or 12th rib (although the zone of apposition does not reach the 12th rib), we performed three experiments in which the pleural liquid pressure was measured in the 6th and 9th ics at the same height in the supine posture. While pleural liquid pressure, averaged over the breathing cycle, was 6–7 cm H2 O greater in apposition than in lung zone during spontaneous breathing, after passive lung inflation to ∼10 cm H2 O (either in the living paralyzed animal or some minutes post-mortem) pleural liquid pressure was essentially the same in both zones, namely about nil. Hence, the slow liquid flow from apposition to lung zone, which probably occurs during spontaneous breathing (Agostoni et al., 1989), no longer occurs after passive lung inflation. Therefore, the cranio-caudal shear flow, described by the first consideration, takes place unopposed. In order to test whether the above hypothesis is correct we took advantage of the fact that pleural pressure at mid-height of the zone of apposition in supine rabbits is not subatmospheric, and a sample of pleural liquid may, therefore, be taken without passive lung inflation. Hence, in five rabbits samples of pleural liquid in the 9th ics were also taken before passive lung inflation to ∼10 cm H2 O. The labeled albumin concentration of these five rabbits after lung inflation (107 ± 12.4 g/ml) was 50% greater (P < 0.05) than before lung inflation (71.3 ± 7.5 g/ml). This finding supports our hypothesis that passive lung inflation displaces some pleural liquid cranio-caudally. Because of this displacement of pleural liquid, the labeled albumin concentration measured after lung inflation is overestimated in sites caudal to the injection point, and underestimated in sites cranial to the injection point. As a consequence, because without passive lung inflation labeled albumin concentration in the 5th and 7th ics should be similar, the concentration in these, and in the 8th and 9th ics 10 min after the injection might roughly be 90, 90, 80, and 70 g/ml, respectively, instead of
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the values reported in Table 1. The concentration in the other sites should be affected to a minor extent by the phenomenon described above. Even taking this artifact into account, there is a marked difference in concentration between the sites close to and far from injection point. Thirty minutes after the injection the labeled albumin concentration in sites far from the injection point (2nd ics, cranial and caudal mediastinum) was still significantly smaller than in sites close to the injection point (7th and 8th ics). This difference, however, was less than half of the difference between the same sites 10 min after the injection (Table 1). Fifty minutes after the injection, the concentration of labeled albumin in sites far from the injection point was similar to that occurring after 30 min, while the concentration in sites close to the injection point was significantly smaller than that occurring after 30 min. Fifty minutes after the injection the concentration in the 2nd ics was no longer significantly smaller than in sites close to the injection point, while the concentration in the mediastinal sites was still significantly smaller than in sites close to the injection point. This difference, however, was ∼half of the difference between the same sites 30 min after the injection (Table 1). In order to determine the overestimate of the concentration caused by passive lung inflation (see above) after 50 min, we measured the concentration in the 9th ics of four rabbits before and after passive lung inflation. The concentration after lung inflation was 21% greater than before lung inflation, but this difference was not significant. Hence, without lung inflation the concentration in the 7th and 8th ics, 50 min after the injection might roughly be 60 g/ml, i.e., close to the concentration in sites far from the injection point (Table 1). Even after 50 min the distribution of concentration does not seem to be affected by the number of deep breaths (4–13) occurring during the experiments, ventilation or sampling sequence. The time course of the labeled albumin concentration in sites far from the injection point can be obtained from the data in Table 1, taking into account that the initial concentration is zero, and probably remains so during the first few minutes (Fig. 2, continuous line). The time course of the concentration in a site close to the injection point can also be drawn (Fig. 2, broken line). In this connection, it should be remembered that the labeled albumin concentration in the latter site
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Fig. 2. Time course of concentration of labeled albumin in pleural liquid of various sites after injection of a 0.1 ml bolus with tracer through 6th ics. Continuous line: concentration in sites far from injection point (2nd ics, cranial and caudal mediastinum); initial concentration was zero, and likely remained such during first few min. Broken line: concentration close to injection point (7th ics). Arrow indicates estimated mixed, initial concentration (see text).
is somewhat overestimated (see above). It, therefore, appears that with such a small bolus, mixing is essentially reached after ∼1 h (at least for the requirement of the second aim of this research, see Sections 1 and 4.2). 3.2. Bolus of 0.5 ml for 10 min and of 1 ml for 30 min The labeled albumin concentration in various sites of the pleural space 10 min after the injection through
the 6th ics of 0.5 ml of Ringer-albumin with 500 g/ml of labeled albumin is reported in Table 2. The concentration in sites far from injection point (2nd ics, cranial and caudal mediastinum) was significantly smaller than in sites close to the injection point (7th and 8th ics). The labeled albumin concentration in various sites of the pleural space 30 min after the injection through the 6th ics of 1 ml of Ringer-albumin with 500 g/ml of labeled albumin is also reported in Table 2. The concentration in sites far from injection point was not significantly different from that in sites close to the injection point. Hence, with a 1 ml bolus (0.38 ml/kg) mixing is essentially reached after 30 min, while with a 0.1 ml bolus it is reached after ∼1 h (see Section 3.1). Though the labeled albumin concentration was the same in the boluses of different volumes, the labeled albumin concentration in the pleural liquid is greater when the bolus injected is larger, because of the smaller dilution of the bolus by the physiological pleural liquid. Therefore, in order to compare the distribution of the labeled albumin concentration between boluses of different volume, the concentration must be expressed as a per cent of the corresponding Cmix (see Sections 2.1 and 3.1). Fig. 3 provides a synoptic view of the distribution of the labeled albumin concentration expressed . When analyzing this histogram, it should as % Cmix be remembered that the concentration in the 7th and 8th ics is somewhat overestimated because of the artifact produced by passive lung inflation, and that this phenomenon tends to vanish when the differences in concentration are small. It appears from this histogram that when the volume of the bolus is increased from 0.1 to 0.5 ml (10 min in both cases) the concentration distribution becomes more uniform, nearly as much as when
Table 2 Labeled albumin concentration (g/ml) in various sites of the pleural space 10 or 30 min after injection through the 6th intercostal space of 0.5 or 1 ml of Ringer-albumin, respectively, with 500 g/ml of labeled albumin Bolus volume (ml)
0.5 1.0
Time (min)
10 30
n
9b 5
Body weight (kg)
2.64 ± 0.11 2.65 ± 0.16
Estimateda in mixed, initial pleural liquid
Sampling site Intercostal space
Mediastinum
2nd Cranial lobe
7th Caudal lobe
8th Caudal border
Cranial Middle or cranial lobe
Caudal Accessory lobe
166 ± 16.9 302 ± 30.1
299 ± 13.4 310 ± 7.6
298 ± 12.2 331 ± 7.2
145 ± 26.4 308 ± 43.7
188 ± 31.4 290 ± 27.3
Values are mean ± S.E. a See text. b Except cranial mediastinum in which n was 7.
234 ± 3.1 318 ± 4.6
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Fig. 3. Concentration of labeled albumin in pleural liquid, expressed as per cent of estimated mixed, initial concentration (see Sections 2.1 and 3.1), at various times and with boluses of different volume. Concentration in 7th and 8th ics is somewhat overestimated (particularly at 10 min) because of passive lung inflation (see Section 3.1).
the time is increased from 10 to 30 min with the bolus of 0.1 ml. Moreover, when the volume of the bolus is increased from 0.1 to 1 ml (30 min in both cases), the concentration distribution becomes more uniform than when the 0.1 ml bolus is kept for 50 min.
4. Discussion 4.1. Bolus distribution The results show that in supine rabbits a small liquid bolus (0.04 ml/kg) injected into the pleural space through the 6th intercostal space (ics) does not preferentially reach the mediastinal regions, as it has been maintained for supine dogs on the basis of the images taken by a gamma camera placed horizontally over the chest, thus providing the frontal projection of the activity of 99m Tc-albumin injected with a small bolus (0.05–0.08 ml/kg) into the pleural space through the 6th ics (Miserocchi et al., 1984; Negrini et al., 1985). Indeed, in our experiments the labeled albumin concentration in mediastinal regions, 50 min after the injection, was not greater than in the regions close to the injection point, even taking into account the overestimation of the concentration in the 7th and 8th ics caused
by passive lung inflation (Table 1 and Fig. 2). This discrepancy might be due to a species difference, but this does not seem to be the case, because even during the first few minutes the distribution of a dye injected with a small bolus through the 6th ics space of dogs is different from that obtained with a frontal projection of the activity of radiolabeled albumin (see Section 1). It, therefore, seems likely that the discrepancy between the present data on rabbits and those obtained in dogs with radiolabeled albumin depends on the deceptive image provided by the frontal projection of the activity of the tracer. Indeed, the following points should be taken into account, as partially done by the authors (Miserocchi et al., 1984; Negrini et al., 1985). (1) For a given concentration of tracer per unit of pleural surface and a given distance from the scanning plane, the regional signal on the plane increases with the increase in the angle formed by that area of pleura with the scanning plane. (2) Radiation energy per unit surface of the scanning plane (and hence, signal intensity) decreases markedly with the distance between the plane and the source of radiation. Moreover, this attenuation is enhanced by the density of the tissue crossed by the radiation. (3) Signal intensity depends on the sum of the frontal projections of all the regions of pleura at various depths subtended by a given area of the scanning plane. In
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addition, the initial part of the lymphatics draining the pleural space are in the field scanned by the gamma camera. Therefore, the complex geometry of the pleural space in some regions combined with the above three points on radiation measurement defies an analysis of tracer concentration in various regions of the pleural space based on a projection of tracer activity. This conclusion is supported by the following consideration. According to the image provided by the frontal projection on the gamma camera, the specific activity of the tracer in the mediastinal region, and in the latero-diaphragmatic lung region (another region preferentially reached by the bolus according to the authors), 1 h after the injection is three–four times greater than in the other regions of the pleural space (including those surrounding the injection point). This situation persists throughout the next hour (see Fig. 4 in Negrini et al., 1985), during which there is only a slow decay of signal intensity in all regions (varying in rate among them), likely related to the removal of the tracer from the pleural space, and its dilution by the liquid filtering into the pleural space. According to the authors, these marked differences in labeled albumin concentration are due to the preferential movement of the bolus to regions where pleural liquid pressure at a given height is more subatmospheric (mediastinal and diaphragmatic regions). On the other hand, the persistence of the marked differences in signal from the various regions, despite breathing and cardiac movements, suggests that they do not depend on differences in tracer concentration in the various pleural regions, but mainly on the factors mentioned in the above three points on radiation measurement. Indeed, our results show that 50 min after the injection into the pleural space of a small bolus, the labeled albumin concentration only differs slightly among various regions of the pleural space (see Section 3.1). The study of the distribution in the pleural space of a small bolus with radiolabeled albumin (Miserocchi et al., 1984) was mainly intended to support the finding by the same authors that pleural liquid pressure in the mediastinal regions is more subatmospheric than in the costal region at the same height (Miserocchi et al., 1981, 1984). Indeed, this difference in pressure was questionable, because the lower pressure in the mediastinal region might be due to the distortion caused by the introduction of the cannula in this region (Agostoni and D’Angelo, 1991). The present finding
that a small bolus does not preferentially reach the mediastinal region casts further doubts on the more subatmospheric pressure in the mediastinal region. At this stage, it should be borne in mind that the topographical distribution of the bolus changes with the position of the point of its injection, and that our data only refer to injection through the 6th ics. For instance, if the bolus were injected into the 3rd ics, the labeled albumin concentration after 10 min should be greater in the 2nd ics and the cranial mediastinum than in the 7th, 8th ics, and the caudal mediastinum. Moreover, each approach has its own limitations. Our approach does not provide information on the labeled albumin concentration in the interlobar and diaphragmatic lung regions; this limitation, however, should not prevent the information required by the purposes of the present research from being obtained. Sampling in the mediastinal regions takes ∼14 min after the death of the animal, but this delay does not seem to appreciably affect the labeled albumin concentration (see Sections 2.1 and 3.1). Finally, our approach requires passive lung inflation for sampling (see Section 2.1), except for the zone of apposition between diaphragm and rib cage where, in rabbits, pleural pressure is not subatmospheric. Passive lung inflation seems to displace some pleural liquid cranio-caudally (see Section 3.1). This liquid displacement increases the concentration of labeled albumin in the zone of apposition. On the basis of this increase, an approximate correction of the concentration in the regions surrounding the injection point was made (see Section 3.1). Unfortunately, this effect of passive lung inflation was only measurable in one site. 4.2. Mixing of the bolus and removal of labeled albumin Complete mixing of a small bolus is never reached because after 30 min the quantity of labeled albumin removed from the pleural space becomes appreciable, and the overall concentration, therefore, decreases because of liquid filtration into the pleural space when the bolus is small (Bodega and Agostoni, 2004). As a consequence, the concentration in sites far from the injection point no longer increases. Another factor preventing a complete mixing is the uneven removal of labeled albumin produced by the lymphatic drainage through the stomata, which are unevenly distributed
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(Wang, 1975, 1985; Negrini et al., 1991). This way of removal alone does not change the labeled albumin concentration, but since some liquid is filtered into the pleural space, the labeled albumin concentration decreases. This decrease in concentration should become greater from the 2nd to the 7th and 8th ics, and from these to the mediastinal regions, because the stomata are lacking in the apical region, and are more concentrated in the mediastinal than in the middle intercostal region. This phenomenon does not appear before 30 min because the prevailing changes in concentration of labeled albumin during this period are those due to the mixing of the liquid of the regions close to the injection point with liquid of the other regions. After 30 min, these changes become smaller, and therefore, those due to the uneven distribution of the stomata become apparent. The contribution of the lymphatic drainage through the stomata of the parietal mesothelium to the overall removal of labeled albumin from the pleural space has recently been measured in anesthetized rabbits with a small (0.3 ml, 0.12 ml/kg) or large (6 ml, 2.4 ml/kg) hydrothorax of Ringer-albumin with labeled albumin and labeled dextran-2000 kDa (Bodega and Agostoni, 2004). This dextran was used as marker for liquid removal through the stomata because, owing to its size, it should essentially leave the pleural space only through the stomata. The quantity of labeled albumin removed by the lymphatic drainage through the stomata was determined from the product of the volume of liquid removed through them and the estimated mean concentration of labeled albumin in the pleural liquid during the experiment. The removal of labeled albumin through the stomata turned out to be 39% of the overall removal in the small hydrothoraces and 64% in the large ones. On the other hand, it could be argued that the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin in the small hydrothorax has been substantially underestimated because of the following considerations. (1) The distribution of stomata in the parietal pleura is quite uneven (see above), (2) the above experiments lasted 3 h, and (3) the mixing time of a small hydrothorax was unknown. The assessment in the present research of the time required for approximate mixing of small and medium boluses (injected through the 6th ics as in the previous research; Bodega and Agostoni, 2004) provided the information for an analysis of this prob-
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lem. The mixing times for boluses of 0.1, 0.5, and 1 ml are ∼1, 3/4, and 1/2 h, respectively (see Section 3.1). Hence, for the small hydrothorax of the previous research (0.3 ml) the mixing time might have been nearly 1 h, i.e., substantial compared with the duration of the experiment (3 h). Instead, for the large hydrothorax (6 ml) the mixing time might have been less than 10 min, i.e., almost negligible. Hence, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin might have been appreciably underestimated in the small, but not in the large hydrothorax. Nevertheless, taking into account that the 6th ics is about equidistant from the region without stomata (apical region) and those with the highest concentration of stomata (mediastinal and diaphragmatic ones) (Wang, 1975, 1985; Negrini et al., 1991), the above underestimation might be small. The removal of labeled albumin through the stomata in the small hydrothorax was 12.6 g in 3 h, which corresponds to 39% of the overall removal (31.9 g; Bodega and Agostoni, 2004). Because mixing time in this hydrothorax is somewhat less than 1 h (see above), the removal of labeled albumin through the stomata during the 2nd and 3rd hour should not be affected by this phenomenon, while removal during the 1st hour could be smaller because of the uneven distribution of labeled albumin combined with the uneven distribution of stomata. To be safe, in the following example, the removal of labeled albumin through the stomata during the 1st hour has been taken as ∼half of that occurring with uniform distribution of labeled albumin. Therefore, an approximate estimate of the removal of labeled albumin through the stomata during each of the 3 h may be 2.6 g for the 1st and 5 g for the 2nd and 3rd, i.e., 12.6 g for the whole period (for simplicity, we disregarded the small difference in removal among hours caused by the 15% decrease in labeled albumin concentration during the experiment; Bodega and Agostoni, 2004). Instead, in case of an instantaneous mixing, the removal of labeled albumin through the stomata during the 1st hour would be greater, being similar to the removal during the 2nd or 3rd hour, and therefore, the removal during the whole 3-h period would also be greater than that found experimentally (12.6 g). Hence, in this virtual case an estimate of the removal of labeled albumin through the stomata during each of the 3 h would be 5 g, i.e., 15 g for the whole period. That is, the uneven distribution of
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labeled albumin during the mixing time, combined with the uneven distribution of the stomata, should involve a smaller removal of labeled albumin through the stomata of 2.4 g. The loss of labeled albumin through the other mechanisms (transcytosis, convection and diffusion) is 19.3 g, being given by the overall removal (31.9 g) minus the removal through the stomata (12.6 g). Assuming that the loss of labeled albumin through the other mechanisms did not change because of the relatively long mixing time, the overall removal of labeled albumin in case of instantaneous mixing would be 15 + 19.3 = 34.3 g, instead of 31.9 g (see above). Hence, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin would be of 15 over 34.3 g, i.e., 44% instead of 39%. Therefore, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin from the small hydrothorax would have been underestimated by only 5%. Furthermore, if the removal of labeled albumin through the other mechanisms also decreased because of the relatively long mixing time of the small hydrothorax, with instantaneous mixing this removal would be greater, and therefore, the overall removal of labeled albumin would also be greater. As a consequence, the contribution of the lymphatic drainage through the stomata to the overall removal of labeled albumin would be less than 44%. That is, the underestimate would be less than 5%. The above analysis, therefore, supports the conclusion of the previous research (Bodega and Agostoni, 2004), namely that under physiological conditions (i.e., with a volume of liquid in the pleural space even smaller than that with the small hydrothorax) the lymphatic drainage through the stomata does not contribute most of protein and liquid removal from the pleural space, contrary to what has previously been maintained by others (for literature see Bodega and Agostoni, 2004). 4.3. Pleural liquid thickness The indirect evidence provided by the present research that passive lung inflation displaces some pleural liquid cranio-caudally suggests to reconsider the interpretation by Wang and Lai-Fook (1997) of their findings that pleural liquid thickness increases from the 2nd to the 8th ics in rabbits ventilated by positive pressure. To this end, one has first to recall a previous research by the same authors in which the thickness
of pleural liquid of rabbits ventilated with positive pressure was found to increase with the increase in ventilation (Wang and Lai-Fook, 1993). The pleural liquid thickness was measured by fluorescence videomicroscopy through a parietal pleura “window” in the 4th or 6th ics. According to their interpretation, the increase in pleural liquid thickness with the increase in ventilation is due to liquid displacement from the lobar margins, which would decrease power dissipation when the velocity of lung movements is increased (Wang and Lai-Fook, 1997). On the other hand, the above finding could simply be the consequence of positive pressure ventilation, which by increasing the pressure on the pleural surface displaces some pleural liquid beneath the “window”, which is more compliant than the rest of the chest wall (Agostoni and Zocchi, 1998). Indeed, during positive pressure ventilation the thickness of the pleural liquid beneath the “window” increased at each lung inflation (see Fig. 1 of Wang and Lai-Fook, 1993). In the following research on rabbits ventilated by positive pressure, in addition to the cranio-caudal increase in the pleural liquid thickness mentioned above, Wang and Lai-Fook (1997) found that a marked increase in ventilation did not increase the pleural liquid thickness in the 2nd ics, but did it moderately in the 5th ics, and markedly in the 8th ics. Because the relative velocity of the lung increases in the cranio-caudal direction for a given ventilation, they concluded that the increase in pleural liquid thickness from the cranial to the caudal regions has a functional relevance because it provides a uniform shear stress despite the different velocity of the lung in these regions. On the other hand, as pointed out by the authors, pleural liquid thickness is not a function of lung velocity alone, because with two values of ventilation the pleural liquid thickness was smaller in the 5th ics than in the 8th ics despite similar lung velocity. In connection with these findings and their interpretation the following considerations may be made. (1) The present research provides indirect evidence that passive lung inflation displaces some pleural liquid cranio-caudally. Therefore, during positive pressure breathing, pleural liquid thickness in the caudal ics should increase at each lung inflation. (2) The above-mentioned displacement of pleural liquid under the “window” caused by positive pressure breathing might be greater in the caudal than in the cranial “window” because the caudal ics are likely more
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compliant than the cranial ones because of the greater distance between the ribs. This may further contribute to the increase in pleural liquid thickness in the caudal “window” occurring during positive pressure breathing. (3) The increase in pleural liquid thickness with the increase in lung velocity would have a functional relevance if pleura lubrication were hydrodynamic, but a recent research has shown that it is boundary, i.e., frictional force is independent of velocity (D’Angelo et al., 2004). Therefore, an increase in thickness of the pleural liquid with the increase in lung velocity would not be of functional relevance.
Acknowledgment We thank R. Galli for skillful technical assistance.
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