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1 Trauma Research Program and Department of Laboratory Medicine and Pathobiology, Sunnybrook and Women's College Health Sciences Centre, University of Toronto, Toronto, Ontario, Canada M4N 3M5; and 2 Department of Medicine, University of Rochester, Rochester, New York 14642
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ABSTRACT |
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 Top
 Abstract
 Introduction
Materials and methods
Results
Discussion
References
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The objective of this study was to determine the relative roles of arachnoid villi and cervical lymphatics in the clearance of a cerebrospinal fluid (CSF) tracer in rats. 125I-labeled human serum albumin (125I-HSA; 100 µg) was injected into one lateral ventricle, and an Evans blue dye-rat protein complex was injected intravenously. Arterial blood was sampled for 3 h. Immediately after this, multiple cervical vessels were ligated in the same animals, and plasma recoveries were monitored for a further 3 h after the intracerebroventricular injection of 100 µg 131I-HSA. Tracer recovery in plasma at 3 h averaged (%injected dose) 0.697 ± 0.042 before lymphatic ligation and dropped significantly to 0.357 ± 0.060 after ligation. Estimates of the rate constant associated with the transport of the CSF tracer to plasma were also significantly lower after obstruction of cervical lymphatics (from 0.584 ± 0.072/h to 0.217 ± 0.056/h). No significant changes were observed in sham-operated animals. Assuming that the movement of the CSF tracer to plasma in lymph-ligated animals was a result of arachnoid villi clearance, we conclude that arachnoid villi and extracranial lymphatic pathways contributed equally to the clearance of the CSF tracer from the cranial vault.
cerebrospinal fluid; cerebrospinal fluid drainage; rat brain; spinal cord; lymph nodes
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INTRODUCTION |
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Top
Abstract
Introduction
Materials and methods
Results
Discussion
References
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THE DRAINAGE of cerebrospinal fluid (CSF) from the brain plays an important role in maintaining homeostasis within the cranial cavity. It is believed that the anatomic element responsible for transporting CSF from the cranial vault is the arachnoid villus with the pressure difference between the CSF and dural venous sinuses providing the dominant driving force for CSF absorption (reviewed in Ref. 11). Most vascularized tissues contain a lymphatic circulation to remove filtered liquid and protein from the interstitial compartment, but the central nervous system (CNS) is unusual in that it does not contain lymphatic vessels. Nonetheless, CSF and extracranial lymph compartments are linked physiologically, because tracers injected into cranial CSF enter lymphatic vessels in the head and neck region (reviewed in Ref. 7).
Despite the tracer recovery data and anatomic studies in cats (6), rabbits (8, 9, 20), rats (17), and sheep (5) that suggest an important role for lymphatics in the removal of CSF, it has always been assumed that the amount of CSF removed by extracranial lymphatic vessels is very small. Part of the difficulty is that estimation of the absorption of CSF into lymphatics presents several experimental problems. First, from an anatomic perspective, the lymphatic drainage pathways are complex and cannot be studied easily. The lymphatic vessels believed to drain CSF collect lymph not only from the CNS but also from many other tissues and organs. Second, the relative contribution of arachnoid villi and lymphatics to CSF absorption has never been determined; therefore, it was difficult to put the role of lymphatics into perspective. Third, tracer recovery studies can be problematic. The transport of a CSF tracer into plasma by the arachnoid villi would filter back into the lymphatic compartment, resulting in an overestimate of the lymphatic contribution to CSF absorption.
These issues have recently been addressed in studies by our group. We demonstrated that thoracic duct and cervical lymph diversion/ligation in conscious sheep reduced the plasma recovery of intracerebroventricularly injected human serum albumin (HSA) by ~50% (3). We also estimated CSF volumetric clearance via arachnoid villi and lymphatics separately using tracer recovery data. A flow marker (125I-HSA) was infused into the ventricles. A second tracer (131I-HSA) was injected intravenously to calculate loss of the CSF flow marker from the circulation and to estimate the rate of 125I-HSA recirculation from the plasma into the lymphatics. Mass balances were carried out around the plasma, the retropharyngeal-cervical lymph pathway, and the thoracic duct, and the equations were solved for the rates of drainage via lymphatics and arachnoid villi. These experiments demonstrated that extracranial lymphatic vessels transported CSF from the cranial vault in volumes roughly equivalent to those drained through arachnoid villi (4).
The sheep data challenge the central tenet that CSF outflow occurs primarily through arachnoid villi and granulations. However, only in sheep has the quantitative significance of lymphatic CSF drainage been addressed. Quantitation of lymphatic CSF transport in other species needs to be investigated to be certain that an important role for lymphatics in CSF clearance is not a species-specific phenomenon. The rat would appear to be an obvious candidate for this analysis because the anatomic relationships between the CSF and extracranial lymph compartments have been elucidated in this animal. After the injection of india ink into the cisterna magna, carbon particles were observed passing from the subarachnoid space beneath the olfactory bulbs into discrete channels that passed through the cribriform plate into nasal submucosal lymphatic vessels (17). The carbon particles were not observed to spread diffusely through the interstitium of the nasal submucosa, suggesting that CSF transports directly into lymphatic vessels in this species rather than indirectly through an intermediate interstitial compartment, as has been proposed by other investigators (16). As a consequence, cervical lymphatic vessels in the rat may take up CSF more rapidly and absorb a greater proportion of the total CSF volume removed from the cranial vault.
The purpose of the experiments reported here was to investigate the lymphatic and arachnoid villi contribution to the transport of a CSF protein tracer in rats. Specifically, we injected HSA into the CSF compartment and compared its transport to plasma before (sum of arachnoid villi and lymphatic transport) and after (arachnoid clearance only) ligation of the cervical lymphatic vessels in the same animals. The differences in plasma recoveries and in the CSF-to-plasma tracer transport rates in the two phases of the experiment would provide an estimate of the lymphatic contribution to CSF tracer clearance.
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MATERIALS AND METHODS |
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Top
Abstract
Introduction
Materials and methods
Results
Discussion
References
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Surgery
Male Wistar rats weighing 350-650 g (Charles River) were used in this investigation. Experiments were approved by the ethics committee at the Sunnybrook Health Science Centre and conformed to the guidelines set by the Canadian Council on Animal Care and the Animals for Research Act of Ontario.Rats were anesthetized with a mixture of 2.5 mg/kg acepromazine and 80 mg/kg ketamine HCl injected intraperitoneally. Anesthesia was maintained using a 1:1 solution in sterile saline of pentobarbital sodium (65 mg/ml). Surgery was aided with the use of a dissecting microscope (Wild Leitz, Willowdale, ON, Canada). A sagittal incision was made in the rat's scalp to expose the coronal suture. A 25-gauge needle was used to bore a hole 2 mm caudal to the coronal suture and 2 mm lateral to the saggital suture. A brain infusion catheter (Alza, Palo Alto, CA) was introduced into one of the lateral ventricles and secured to the skull with cyanoacrylate glue. Patency was checked by infusing a small amount of sterile saline through the catheter. At the end of the experiment, Evans blue dye was injected into the lateral ventricle to confirm placement of the infusion catheter. For injection of the vascular tracer, a catheter (PE-10 catheter, Clay Adams, Parsippany, NJ) was placed in one of the femoral veins. A femoral artery was cannulated (0.58 × 0.96 mm Silastic tubing, Dow Corning, Midland, MI) to permit sequential blood sampling. Patency of both catheters was maintained using a heparinized saline flush. Both catheters were exteriorized and attached to the hind leg with tape. All skin incisions were closed using 3-0 silk.
Experimental Protocols
Qualitative studies using 125I-HSA. One hundred micrograms of 125I-HSA (0.0093 MBq) was injected into one of the lateral ventricles in each of eight anesthetized rats. Six hours later the animal was killed. Various lymph nodes and selected other tissues present within the limbs, body cavities, and neck of the animal were excised, weighed, and counted for radioactivity. In cases where nodes were bilateral or arranged in chains, a single point was plotted representing the mean radioactivity per gram tissue for that group of nodes. The results of these experiments were calculated as percent injected dose per gram tissue.Quantitative studies. In phase 1 of the experiment, radiolabeled HSA was injected into a lateral ventricle, and the recovery of the tracer was monitored in blood. Transport of the CSF tracer to plasma occurs through both arachnoid villi and extracranial lymphatics. After 3 h the cervical lymphatics were ligated in the same animal, and the experiment was repeated with the same mass of HSA tagged with a different radiolabel. The plasma recovery in phase 2 of the experiment was assumed to represent arachnoid villi transport, because the major CSF-lymph drainage pathways had been interrupted.
In the first phase of the experiment, 100 µg of
125I-HSA (CSF tracer, 0.0093 MBq)
was injected into a lateral ventricle. Evans blue dye was mixed with
plasma from a donor rat of the same strain ex vivo (0.1% in rat
plasma) and was injected into the femoral vein. Blood samples (250 µl) were taken at 10, 14, 18, and 22 min to determine the volume of
distribution of the protein tracer and, after this, hourly for 3 h.
In phase
2 of the experiment ~1 h later, the
lymph nodes in the cervical chain (bilateral posterior cervical,
anterior cervical, and submaxillary lymph nodes) were injected with
Evans blue dye to visualize the lymphatic vessels. All detected vessels
were ligated with 8-0 silk sutures as illustrated in Fig.
1. After this, 100 µg of
131I-HSA (0.37 MBq) was injected
into the lateral ventricle, and blood samples were collected hourly for
three more hours. In sham animals,
phases
1 and
2 of the experiment were identical
except that no lymphatic vessels were identified with Evans blue or
ligated. All blood samples were collected in Microvette tubes
containing EDTA (Sarstedt, St. Leonard, PQ, Canada) and
centrifuged for plasma collection.
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Tracers and Solutions
125I-HSA (0.93 MBq/ml, 10 mg/ml) and 131I-HSA (37 MBq/ml, 10 mg/ml) were obtained from Frosst (Kirkland, PQ, Canada). Lactated Ringer solution was purchased from Baxter (Chicago, IL). Each lot was tested for protein-free radioactivity by passing through a Centricon centrifugal concentrator (10,000 mol wt cutoff). To ensure that the measured radioactivity in any collected sample was protein associated, a second set of aliquots was assayed after precipitation with 10% trichloroacetic acid. Free, or non-protein-associated, 125I or 131I represented <1% of the total radioactivity in any sample.Analysis of Data
We plotted the nondimensional tracer mass vs. time for each experimental condition (cpm × plasma volume/cpm injected) to assess the effects of lymphatic ligation or sham surgery on the plasma recoveries of the CSF tracer. However, as HSA enters the plasma, some of this protein will filter out of the vascular compartment with the result that plasma recoveries will be underestimated. To correct the data for the loss of filtered protein, an elimination rate (Kexp) was defined and entered into a mass balance equation.A two-compartment model (CSF and plasma) was used to quantify rates of
transfer of the tracer from the CSF to the blood. The CSF and plasma
compartments were assumed to be well mixed and constant in volume. The
labeled protein was injected into the CSF compartment and was assumed
to be contained entirely within this compartment at
time
0. It was further assumed that
transfer of the tracer from the CSF to the plasma occurred at a
constant turnover rate
[(KB)(h
1)]
without sequestration in the brain parenchyma or other tissues. A mass
balance around the plasma yields
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(1) |
A mass balance around the CSF compartment yields
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(2) |
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(3) |
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(4) |
The elimination of tracer from plasma was determined from the slope of the plasma disappearance of the Evans blue-protein complex (Kexp). To calculate Kexp, plasma concentrations of Evans blue-protein complex were divided by the initial concentration (C0) for normalization. Kexp was defined as the slope of the log-transformed data. The optical density of Evans blue in each plasma sample (624 nm) was read from a spectrophotometer, and the dye concentrations (from a standard curve) were extrapolated graphically to time 0. The volume of distribution of the tracer in plasma was calculated from the dilution of the Evans blue dye-rat albumin complex. All data were expressed as means ± SE. The results were analyzed with ANOVA or a paired Student's t-test as appropriate. We interpreted P < 0.05 as significant.
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RESULTS |
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Top
Abstract
Introduction
Materials and methods
Results
Discussion
References
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Anatomic Studies With 125I-HSA
As part of their normal physiological function, lymphatic vessels absorb extravascular protein and, after passage through various lymph nodes, return it to the vasculature. Taking advantage of this function, 125I-HSA was injected into a lateral ventricle to help elucidate lymphatic pathways that drain CSF from the brain. Increased radioactivity would indicate the presence of 125I-HSA in transit through the lymphatic channels in the nodes. Additionally, one would expect to see 125I-HSA in the nodal blood because some of the tracer would have exited the CSF via the arachnoid villi. In the highly vascular spleen, for example, we monitored 0.16% injected dose/g tissue (Fig. 2). The less vascular fat and skeletal muscle tissues contained less radioactivity. Lymph nodes that one would not expect to drain CSF (e.g., popliteal and axillary nodes) contained levels of radioactivity similar to those measured in the spleen. All of the nodes tested in the head and neck region had elevated radioactivity compared with the splenic tissues (posterior and anterior cervical nodes as well as the submaxillary nodes). In addition, the radioactivity in the head and neck lymph nodes was significantly greater than that measured in the peripheral popliteal node, which was not expected to be positioned on a CSF drainage pathway. The fact that the radioactivity in the lumbar nodes was significantly greater than that in the spleen or popliteal nodes suggested the transport of the protein tracer from spinal CSF into lymph.
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Recoveries of CSF Tracer and Transport Rate of HSA Into Plasma
Comparisons of the fractional recoveries of intraventricularly injected tracer in plasma over time demonstrated a significant reduction after ligation of the cervical lymphatic vessels (Fig. 3). In contrast, no significant change in the fractional recoveries were observed after sham surgery. These recovery data were not corrected for the loss of tracer from the plasma compartment.
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Estimates of the constant
(KB) that
determined the rate of transport of the CSF tracer to plasma
(calculated from Eq. 4) were consistent with the raw recovery data.
KB values were
significantly lower after the cervical lymphatics were ligated (Fig.
4). Before ligation,
KB averaged 0.584 ± 0.072/h and after ligation dropped to 0.217 ± 0.056/h. In the
sham experiments estimates for
KB increased from
0.601 ± 0.102/h before sham surgery to 0.709 ± 0.144/h in phase
2 of the experiments, but these
effects were not significant. The average animal weights,
plasma volumes, and calculated values for
Kexp are
illustrated in Table 1.
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DISCUSSION |
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Top
Abstract
Introduction
Materials and methods
Results
Discussion
References
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The objective of this study was to compare the relative roles of arachnoid villi and extracranial lymphatics in the clearance of a CSF protein tracer from the rat brain. Cervical ligation reduced the rate constant associated with the transport of the CSF tracer to plasma by >60% and the fractional plasma recoveries of the CSF tracer by ~50%. Therefore, we estimate that at least one-half of the CSF tracer cleared from the cranial vault in rats is removed by cervical lymphatics. In previous studies, we estimated that extracranial lymphatics in sheep transported approximately one-half of a CSF protein tracer to plasma (3). The similarities between the rat and sheep data occur despite the fact that the CSF turnover rate in rats (~2 h, Ref. 1) is faster than that observed in sheep (ranging from 4 to 5 h, Ref. 10). Anatomic factors may influence the flow of CSF in the two species. In sheep the increased cerebral convexities provide a less uniformly distributed CSF pool, and the multiple cisterns may retard directionalized tracer flow. Conversely, in rats the subarachnoid velae provide an early communication between the ventricles and subarachnoid space (13) and could allow rapid removal of CSF. In any event, the proportion of the CSF tracer drained through arachnoid villi and extracranial lymphatic vessels was very similar in both species. This suggests that the distribution of CSF clearance through the two pathways is independent of the CSF turnover rate and that the role of lymphatics in CSF transport is not a species-specific phenomenon. Rather, cervical lymphatics may have an important and consistent role in venting cranial CSF in many if not all mammals.
Methodological Considerations
Mathematical model. The mass balance equations and the generation of the rate coefficient KB was an attempt to provide a more realistic value for the CSF-to-plasma transport of the protein tracer. Because the movement of HSA from the CSF into plasma is accompanied by simultaneous filtration of the protein out of the plasma into the various tissue compartments, plasma recovery data inherently underestimate the transport process. A coefficient of HSA elimination from plasma was defined (Kexp) and incorporated into a mass balance equation designed to correct the plasma data for the tracer loss. With the current assumptions, however, the estimates of KB may be in error depending on unmeasured experimental variables. Nonetheless, the magnitude of the change in KB values associated with obstruction of the cervical lymphatic pathways was very similar to that observed for the uncorrected plasma recoveries of the CSF tracer. Both measures of tracer transport support the major conclusion of the study that extracranial lymphatic vessels remove about one-half of the CSF protein tracer from the cranial vault.Issues related to anatomy of lymphatic CSF drainage pathways. Published accounts suggested that the cervical ducts draining the nasal submucosa were likely the most important CSF-draining lymphatics in the rat (17, 21). Our anatomic studies would support this conclusion. However, it is likely that we underestimated the lymphatic contribution to CSF protein clearance using our experimental approach. First, cervical lymphatics in the rat are very small, and we may have failed to identify all of the relevant vessels. Second, other lymphatics may also contribute to CSF clearance. For example, Kida et al. (17) observed india ink in the dural lymphatics after injection into the cisterna magna. Because of the difficulties associated with the identification and ligation of other potentially relevant lymphatics (e.g., dural vessels and the right lymph duct), the contribution of these lymphatics is unknown. Additionally, the thoracic duct was not ligated, and this vessel may transport CSF tracer from lumbar lymphatics that have absorbed tracer from spinal CSF (18). We were concerned that obstruction of this large lymph trunk would cause a significant physiological disturbance, and because its contribution to CSF drainage from the cranium is probably minor, we decided to leave this vessel intact. These surgical limitations would tend to result in an overestimation of arachnoid villi transport and an underestimation of the lymphatic contribution to the clearance of a CSF protein tracer. We believe that an estimate of 50% of tracer transport via lymphatic pathways represents an average minimum value and that the contribution of extracranial lymphatics to CSF clearance could be much higher.
Another factor to consider is the small (not significant) increase in KB observed between phases 1 and 2 in the sham animals. The increased transport rate in phase 2 was probably due to a rise in intracranial pressure after acute placement of the ventricular catheter (12). An increase in total CSF absorption has been observed with increased pressures (2, 15). However, potential pressure-related factors did not have a significant impact on the results. The plasma recoveries of the CSF tracer in phases 1 and 2 of the sham group were nearly identical (Fig. 3).
Assumptions related to nonlymphatic transport of the CSF tracer. After ligation of the cervical lymphatic vessels, we assumed that plasma recoveries would reflect transport via arachnoid villi, although we have no direct evidence that this is the case. Arachnoid villi in rats are relatively simple structures compared with their counterparts in humans (19). Few arachnoid villi appear to be associated with the superior and inferior sagittal sinuses in this species. Those that are present are situated close to the major nasal lymphatic drainage pathways (17). Additionally, as noted earlier, it is possible that some of the tracer recovered in the plasma entered by lymphatic vessels that were not obstructed. This would mean that estimates for arachnoid villi transport would be inflated by an unknown amount.
Perspectives
Despite the fact that a physiological association between extracellular fluid in the brain and extracranial lymph has been appreciated for over 100 years, this concept has never been embraced by the biomedical community. Because the drainage of CNS interstitial fluid was assumed to occur by clearance through specialized structures termed arachnoid villi, there seemed little need to incorporate lymphatics in the conceptual framework that has driven investigation in this area. The development of methods that allow the separate quantitation of CSF tracer clearance through arachnoid villi and extracranial lymphatic vessels has led to studies that challenge the conventional view. In sheep and in rats, nearly 50% of all CSF tracer drained from the cranial vault is removed by cervical lymphatic vessels. CSF clearance through extracranial lymphatic vessels has important implications. For example, the ease by which CSF is vented from the cranial vault can be estimated as an outflow resistance (reviewed in Ref. 14). The changes in outflow resistance that occur, for example, in some forms of hydrocephalus, subarachnoid hemorrhage, and other pathophysiological states are believed to contribute significantly to elevations of intracranial pressure. It appears likely, however, that a lymphatic component figures prominently in the physiological parameters that define outflow resistance. The question of whether anomalies associated with the cervical lymphatic transport of CSF can contribute to elevations of intracranial pressure would appear to warrant further investigation.| |
ACKNOWLEDGEMENTS |
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The authors thank Ninos Yacoub for technical assistance.
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FOOTNOTES |
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This research was funded by the Medical Research Council of Canada.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests: M. G. Johnston, Trauma Research Program, Dept. of Pathology, Univ. of Toronto, Research Bldg. S-111, Sunnybrook Health Science Centre, 2075 Bayview Ave., Toronto, ON, Canada M4N 3M.
Received 8 July 1998; accepted in final form 18 November 1998.
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Top
Abstract
Introduction
Materials and methods
Results
Discussion
References
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I. Silver, B. Li, J. Szalai, and M. Johnston Relationship between intracranial pressure and cervical lymphatic pressure and flow rates in sheep Am J Physiol Regulatory Integrative Comp Physiol, December 1, 1999; 277(6): R1712 - R1717. [Abstract] [Full Text] [PDF]
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) and after (
) lymph ligation
(n = 7, top). For sham animals
(n = 7, bottom), no significant differences
were observed between first phase of experiment (
) and sham-operated
phase (
).
