The following is the abstract of the article discussed in the subsequent letter.
Hermann, Gerlinda E., R. Alberto Travagli, and Richard C. Rogers. Esophageal-gastric relaxation reflex in rat: dual control of peripheral nitrergic and cholinergic transmission. Am J Physiol Regul Integr Comp Physiol 290: R1570–R1576, 2006; doi:10.1152/ajpregu.00717.2005.—It has long been known that the esophageal distension produced by swallowing elicits a powerful proximal gastric relaxation. Gastroinhibitory control by the esophagus involves neural pathways from esophageal distension-sensitive neurons in the nucleus tractus solitarius centralis (cNTS) with connections to virtually all levels of the dorsal motor nucleus of the vagus (DMV). We have shown recently that cNTS responses are excitatory and primarily involve tyrosine hydroxylase-immunoreactive cells, whereas the DMV response involves both an α1 excitatory and an α2 inhibitory response. In the present study, using an esophageal balloon distension to evoke gastric relaxation (esophageal-gastric reflex; EGR), we investigated the peripheral pharmacological basis responsible for this reflex. Systemic administration of atropine methyl nitrate reduced the amplitude of the gastric relaxation to 52.0 ± 4.4% of the original EGR, whereas NG-nitro-l-arginine methyl ester (l-NAME) reduced it to 26.3 ± 7.2% of the original EGR. Concomitant administration of atropine methyl nitrate and l-NAME reduced the amplitude of the gastric relaxation to 4.0 ± 2.5% of control. This reduction in the amplitude of induced EGR is quite comparable (4.3 ± 2.6%) to that seen when the animal was pretreated with the nicotinic ganglionic blocker hexamethonium. In the presence of bethanechol, the amplitude of the esophageal distension-induced gastric relaxation was increased to 177.0 ± 10.0% of control; administration of l-NAME reduced this amplitude to 19.9 ± 9.5%. Our data provide a clear demonstration that the gastroinhibitory control by the esophagus is mediated via a dual vagal innervation consisting of inhibitory nitrergic and excitatory cholinergic transmission.
To the Editor: Hermann, et al. (3) in the discussion of their paper criticize the paper we recently published in the American Journal of Physiology–Regulatory, Integrative and Comparative Physiology (2). Their first criticism is that we claim to have employed the same reflex stimulating technique as Rogers, et al. (7). As we point out in an earlier exchange of letters to the editor (Am J Physiol Regul Integr Comp Physiol 290: R1151–R1152, 2006), the purpose of our study was to use the same reflex stimulating technique (esophageal distension) but the initial parameter used failed to alter the end point that we were measuring. Using a decrease in intragastric pressure (via a balloon recording) as a marker of gastric relaxation, esophageal distension of 0.2 ml failed to elicit a significant effect on intragastric pressure (see Fig. 1 of Ref. 2) in our experimental preparation. Hence, we had to use a greater volume to evoke a response. A decrease in intragastric pressure was noted in some animals when the volume of distension was 0.6 ml and this decreased further, when the volume was increased in 0.1-ml increments. To elicit stable responses, we used a volume that ranged from 0.6 to 0.8 ml for most of our pharmacological microinjection studies. Our data are comparable to those of Wei, et al. (8) who reported that esophageal distension with 0.5 ml reduced intragastric pressure in the rat. Additionally, in an earlier study of Rogers, et al. (6), a reduction in gastric motility was observed in the rat with a distension volume of 0.4 ml.
The second criticism is that we did not investigate an esophageal-gastric reflex but probably a gastro-gastric reflex. Their reason for assuming this is that the motility traces in Figure 2 of our paper (2) show a transient increase in antral tone and motility during balloon distension. Although there is a sharp gastric contraction at the start of the esophageal distension [see Fig. 2A, Ferreira, et al. (2)], the predominant response was a decrease in intragastric pressure that was comparably long lasting and was always evident before the cessation of the esophageal distension [see Figs. 1 and 2, Ferreira, et al. (2)]. Furthermore, the sharp gastric contraction did not always occur [see trace C of Fig. 2, Ferreira, et al. (2)]; and if it did, it was markedly decreased, and/or completely absent [see, e.g., Figs. 1A, 2C, and 10B, Ferreira, et al. (2)]. It is possible that the transient contraction shown in Figure 2A of our paper (2) is an artifact due to the transient pressure gradient arising from an injection of the fluid used to distend the balloon.
Hermann and colleagues (3) state that in our study we were monitoring antral gastric tone. We disagree; rather we were monitoring global intragastric pressure by an intragastric balloon introduced via the fundus of the stomach and positioned around the corpus/antrum area. The balloon was inflated with 2 to 3 ml of warm saline to produce a baseline pressure of 6 to 15 mmHg. Since the balloon inflation distended approximately the whole stomach, we interpret the resultant baseline pressure as indicative of the global gastric pressure.
As a third criticism of our paper, Hermann, et al. (3) cite the work of Dong et al. (1) to question the appropriateness of our esophageal distension volume. However, Dong et al. (1) did not use the end point of gastric relaxation; instead, their end point was a change in distal esophageal rhythmic contraction.
Furthermore, Hermann, et al. (3) criticize us for using an intragastric balloon inserted via the fundus for measuring gastric tone. We used the same method described by Krowicki, et al. (5) and Krowicki and Hornby (4) to measure gastric tone. In each case, an intraluminal latex balloon was inserted into the stomach through an incision in the fundus for recording intragastric pressure. In our study (2), we did not obtain any evidence for the presence of a gastro-inhibitory nonadrenergic, noncholinergic (NANC) pathway described by Hermann, et al. (3). In the studies of Krowicki and colleagues (4, 5), decreases in intragastric pressure were obtained as evidence for the activation of this pathway. However, failure to perform ipsilateral vagotomy suggests that the effect may be of nucleus tractus solitarius and not of dorsal motor nucleus of the vagus (DMV) origin.
Finally, Hermann, et al. (3) criticize us for using chloral compounds (e.g., chloral hydrate or α-chloralose) as an adjunct to urethane anesthesia. They state that chloral compounds are known to induce adynamic ileus. (We did not use chloral hydrate in our study; only chloralose was used in combination with urethane.) They cite two studies as evidence for adynamic ileus with these compounds (their Refs. 43 and 48). Their Ref. 43 is a paper by Sababi and Nylander and our reading of that paper provides no evidence that chloralose induces any more adynamic ileus than the anesthetic used by Hermann, et al. (3), namely inactin. Their Ref. 48 is a paper of Silverman and Muir in which adynamic ileus was reported for chloral hydrate but not for α-chloralose. In addition, Krowicki, et al. (5) and Krowicki and Hornby (4) used α-chloralose in their studies, which they concluded, provided evidence of a DMV gastric inhibitory vagal pathway, i.e., a NANC pathway.
In conclusion, we affirm the adequacy of our method for recording the end point we were measuring, namely esophageal distension-induced gastric relaxation. Furthermore, it is our assertion that neither the distension volume, nor the anesthetic used in our study interfered in any way with our observations or our conclusions.
- Copyright © 2006 the American Physiological Society
To the editor: We are delighted to see that Gillis and colleagues recognize that our two laboratory groups are not using the same reflex stimulating or recording techniques to study the esophageal-gastric relaxation reflex (see Table 1 and Fig. 1). This was exactly our point in our earlier exchange of Letters to the Editor (Am J Physiol Regul Integr Comp Physiol 290: R1151–R1152, 2006). Unfortunately, this controversy was started in a previous paper from Gillis' laboratory (1) where the authors stated the express purpose of their studies. To quote their paper, they state: “The specific purpose of the present study was to employ the same [emphasis, mine] reflex-stimulating technique as Rogers et al. [Am J Physiol Regul Integr Comp Physiol 285: R479–R489, 2003]. …”
One reason for the apparent differences in the sensitivity of our measurements may have to do with our basic measurement techniques. The strain gauge elements we use can detect luminal stretch forces as small as 75 mg and as large as 5 g. Parallel measurements in which we place strain gauges on the lumen while measuring gastric pressure changes in response to gradual fluid filling shows that this translates into the ability to detect pressure changes smaller than 1 mm and larger than 8 mmHg, as measured with a P75 low pressure transducer. Although not described in their most recent study (1), previous studies performed in the Gillis lab cite the use of Statham P23 transducers to detect intraluminal pressure changes. These transducers have a very broad range (−30 to 300 mmHg) and proportionally lower sensitivity to small changes in pressure. Indeed, many years ago we attempted to use the same transducers without success. This was the main reason we made the change to strain gauge measurements or measurements with pressure transducers designed for low pressure.
While we have tried to offer potential explanations for our very divergent observations, we must leave it that our stimulation and response recording techniques are vastly different than those used by Gillis' laboratory. As such, we were quite successful in observing that esophageal distension elicited gastric relaxation reflexes (2–5 mmHg) at noticeably lower levels of stimulation than those required in the studies by Gillis's laboratory (1). This rather delicate esophageal-gastric reflex was susceptible to pharmacological investigation as we reported in our recent manuscripts (2, 3).
To the Editor: We appreciate Dr. Weschler's interest in our paper (2) and gladly take the opportunity to answer the two comments Dr. Weschler gave in her Letter to the Editor.
First, the question of whether Heer et al.'s (1) results are compatible 1) with osmotically inactive sodium storage, 2) with sodium-potassium exchange, or 3) with a combination of both, can only be answered unequivocally by data on potassium balance. We do hope that it might be possible for Dr. Heer and her coworkers to provide these crucial data.
Second, we are completely aware that it is inappropriate to directly compare the results of our study with those of Heer et al. (1) for several reasons. The most important reason, as we had already pointed out in our paper (2), is that there are striking species differences between dogs and humans with regard to kinetics of Na+ homeostasis and the response to changes in Na+ intake.
In our studies in freely moving dogs, we induced alterations in total body sodium (TBSodium) that covered the range from moderate deficit to large surplus. These alterations in TBSodium were induced by a variety of methods (10 protocols), not just by changing Na+ intake, because it is well known that the effects of changes in Na+ intake on TBSodium are usually very small in normal dogs and rats, as opposed to human beings (for references, see Ref. 2). Furthermore, it is important to remark that all data on changes of TBSodium, total body potassium (TBPotassium), and total body water (TBWater) reported in our paper are not only based on excretion data, but on balance data for all three variables, i.e., Na+, K+, and water. It is most noteworthy in this context that the daily intake of Na+, K+, and water of our dogs was fixed on a per kilogram body mass basis and controlled for its completeness.
The results obtained are a bit more comprehensive than described in Dr. Weschler's letter: we did not only report instances where TBSodium increase was accompanied by a TBPotassium decrease. In fact, we found that primary changes in TBSodium were accompanied by changes in TBPotassium in the majority of protocols. Four scenarios were observed: 1) TBSodium increase accompanied by a TBPotassium decrease, 2) TBSodium increase accompanied by a TBPotassium increase, 3) TBSodium decrease accompanied by a TBPotassium increase, and 4) TBSodium decrease accompanied by a TBPotassium decrease. Most remarkably, the sum of changes in TBSodium and TBPotassium was always accompanied by osmotically adequate changes in TBWater, regardless of the degree and direction of changes of TBSodium and TBPotassium. Accordingly, plasma osmolality remained unchanged in all instances.
Thus, our present results corroborate various previous observations (for references, see Ref. 2) indicating that primary changes in TBSodium are very often accompanied by changes in TBPotassium and that osmocontrol effectively adjusts TBWater to the body's present content of the major cations, Na+ and K+. This is the reason behind the finding that individual changes in TBWater were a markedly stronger function of simultaneous changes in both TBSodium and TBPotassium (R2 = 0.93) than of changes in TBSodium alone (R2 = 0.83).
Therefore, we completely agree with Dr. Weschler's comment that to conclude the Na+ of a positive Na+ balance has been rendered osmotically inactive requires not only that there be no increase in TBWater or osmolality but also a zero potassium balance. To be more precise, the change in TBSodium plus the change in TBPotassium must exceed the total that can be accounted for by changes in TBWater and osmolality.
The published data of Heer et al.'s study (1) include balance data, i.e., data on intake, extrarenal loss, and urinary excretion for Na+ and water. With regard to K+, only urinary excretion data were included (given as μeq/min), i.e., the crucial information whether or not K+ intake was measured or controlled for was not included. It is also not mentioned whether extrarenal K+ loss or its changes with varying Na+ intake were assessed. Thus, the comment made in our paper (2) regarding Heer's study, “The data of Heer's study in humans that hitherto appeared to demonstrate that osmotically inactive Na+ storage is a rapid process, can no longer be regarded as positive proof (sic) for this storage, because K+ balances were not assessed.” was well founded.
It would be most fortunate if Dr. Heer and her coworkers could provide these crucial data and thereby answer the question whether increasing Na+ intake in her subjects was accompanied by 1) osmotically inactive Na+ storage, 2) Na+/K+ exchange, or 3) a combination hereof.
With regard to compartmental redistributions of Na+ and K+, our exemplary calculations are also based on the respective balance data in conjunction with data on plasma Na+ and K+ concentrations. These calculations revealed that, at least in four of our protocols (2 with increase in TBSodium and 2 with decrease in TBSodium) primary changes in TBSodium included redistribution of substantial amounts of Na+ and K+ between extracellular and cellular space. In each case, an (almost) quantitative, osmotically neutral Na+/K+ exchange between the fluid compartments must have occurred. Because this redistribution was observed even with moderate TBSodium changes and occurred rather rapidly and because Na+ moved into cells in two protocols and out of cells in two others, we conclude that cells may serve as a readily available Na+ store. This Na+storage would be osmotically active, as osmotical equilibration is achieved by opposite changes in cellular K+ content.
Considering these results in conjunction with 1) the well-known fact that primary changes in TBPotassium are almost regularly accompanied by compartmental redistribution of Na+ and K+ and 2) some early reports that also found that primary changes of TBSodium can be accompanied by such redistributions (for references, see Ref. 2), it appears conceivable that redistributions could also have occurred in Heer's subjects. However, a quantitative analysis as exemplified in our article's appendix would require a complete set of data, including data on K+ balance.
In her second comment, Dr. Weschler implies that we had directly compared our data with those of Heer et al's study (1). This is not the case; in fact, we clearly pointed out that major differences of our study and those of Titze et al. (3) and Heer et al. (1) include, but are not limited to, study duration and species differences. As already mentioned in our paper, kinetics of Na+ homeostasis and the response to changes in Na+ intake vary considerably among species (for references, see Ref. 2).
Considering a variety of other well-known differences between dogs and humans relevant for Na+, K+, and water homeostasis, for instance regarding 1) extrarenal loss via sweat (dogs only have sweat glands on their paws); 2) maximum urine concentrating ability (about twice as high in dogs as in humans); 3) metabolic turnover (basal turnover rate per kilogram body mass is about twice as high in dogs as in humans); 4) strikingly different feeding behavior of carnivores (as dogs are) and human beings with regard to time courses and amounts of food intake, as well as salt and water intake, we would refrain from comparing intake data between these species on a per kilogram body mass basis.
The results of our balance studies (2) clearly indicate that changes in TBSodium are often accompanied by TBPotassium and frequently include osmotically active Na+/K+ redistributions among fluid compartments, whereas we did not observe osmotically inactive Na+ storage within our 4-day study period in dogs. Thus, with regard to the validity of the notion that, during Na+ retention, large portions of Na+ are usually stored in an osmotically inactive form, we would like to repeat the conclusion expressed in our paper (2): “Further studies are needed that address the time course of TBSodium changes, involve different species, and must include measurements of TBPotassium or its changes.”