Assessment of transcapillary glucose exchange in human skeletal muscle and adipose tissue
Werner Regittnig,1
Martin Ellmerer,3
Günter Fauler,2
Gerald Sendlhofer,3
Zlatko Trajanoski,1
Hans-Jörg Leis,2
Lukas Schaupp,3
Paul Wach,1 and
Thomas R. Pieber3
1Department of Biophysics, Institute of
Biomedical Engineering, Graz University of Technology; and Departments of
2Biochemical Analysis and Mass Spectrometry and
3Internal Medicine, Diabetes and Metabolism, Karl
Franzens University Graz, A-8010 Graz, Austria
Submitted 9 August 2002
; accepted in final form 5 April 2003
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ABSTRACT
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We studied the kinetics of glucose exchange between plasma and interstitial
fluid (ISF) in human skeletal muscle and adipose tissue under fasting
conditions. Five normal human subjects received an intravenous
[6,6-2H2]glucose infusion in a prime-continuous fashion.
During the tracer infusion, the open-flow microperfusion technique was
employed to frequently sample ISF from quadriceps muscle and subcutaneous
adipose tissue. The tracer glucose kinetics observed in muscle and adipose
tissue ISF were found to be well described by a capillary-tissue exchange
model. As a measure of transcapillary glucose exchange efficiency, the 95%
equilibrium time was calculated from the identified model parameters. This
time constant was similar for skeletal muscle and adipose tissue (28.6
± 3.2 vs. 26.8 ± 3.6 min; P = 0.60). Furthermore, we
found that the (total) interstitial glucose concentration was significantly
lower (P < 0.01) in muscle (3.32 ± 0.46 mmol/l) and adipose
tissue (3.51 ± 0.17 mmol/l) compared with arterialized plasma levels
(5.56 ± 0.13 mmol/l). Thus the observed gradients and dynamic
relationships between plasma and ISF glucose in muscle and adipose tissue
provide evidence that transcapillary exchange of glucose is limited in these
two tissues under fasting conditions.
open-flow microperfusion; interstitial fluid; capillary-tissue exchange model
STUDIES INVESTIGATING THE MECHANISMS of transcapillary fluid and
solute transport suggest that, in most vascular beds, small hydrophilic
substances (e.g., glucose) cross the capillary wall primarily by simple
diffusion (9,
27). Thus, for small
hydrophilic solutes, the capillary wall may be regarded as a porous membrane
through which passive solute exchange between the plasma and the interstitial
fluid (ISF) occurs (9,
27,
33). The permeability of
capillary walls to small hydrophilic solutes has been shown to vary
considerably from one microvascular bed to another
(9). These variations in
capillary permeability are likely to be related to variations in the
ultrastructure of the capillary walls in the microvascular beds (e.g.,
fenestrated vs. continuous capillaries)
(46). Because diffusion of
solutes into or out of blood also depends on the time that blood spends in the
capillary, solute exchange across capillary walls is influenced by blood flow
(9,
38). The magnitude of
transcapillary exchange of very small solutes with high capillary permeability
(e.g., potassium in heart) was shown to depend largely on blood supply (i.e.,
flow-limited exchange) (9). But
in the case of solutes with high molecular weight and low capillary
permeability (e.g., inulin in heart), solute exchange was found to be
practically unaffected by blood flow (i.e., permeability-limited exchange)
(9). Therefore, transcapillary
exchange of hydrophilic solutes of the size of glucose may be best described
as intermediate between the two extreme situations of flow-limited and
permeability-limited exchanges
(9,
38). Thus, besides the general
tissue geometry (e.g., capillary surface area), the main factors determining
the transcapillary exchange of a small hydrophilic solute of the size of
glucose seem to be the prevailing blood flow and the capillary permeability to
the solute.
In recent years, the ISF of peripheral tissues has received considerable
attention as an alternative site for the measurement of glucose levels.
Various methods, such as microdialysis
(4,
18), open-flow microperfusion
(40,
51), electrochemical sensors
(22), transdermal extraction
(23,
49), and hypodermic needles
(2,
44), have been proposed for
the assessment of glucose in the ISF rather than in blood. The ISF glucose
values obtained by such methods may then be used to estimate blood glucose
levels. In this way, the ISF-based glucose assessment could potentially
replace the more invasive blood glucose measurements
(35). Ideally, if
transcapillary exchange of glucose were very rapid and unimpeded, the blood
and ISF glucose would be in instantaneous equilibrium so that, irrespective of
the metabolic rates of cell glucose uptake, the ISF glucose concentrations
would be identical to the blood levels at all times. In this ideal situation,
the quality in estimating blood glucose concentrations from ISF glucose values
would depend solely on the degree of precision and accuracy of the ISF glucose
measurements (1). However,
under the circumstances of limited transcapillary exchange of glucose,
transient differences between ISF glucose levels and changing blood glucose
concentrations may occur, and, if there is an additional glucose consumption
by the tissue cells, a blood-to-ISF concentration difference may also exist
under steady-state conditions
(9,
19,
37,
45). Because of these
differences, accurate calculation of blood glucose concentrations from
interstitial glucose measurements requires one to postulate a mathematical
model of the tissue-specific relationship between blood and ISF glucose levels
under steady-state and non-steady-state conditions. Obviously, in the case of
limited transcapillary glucose exchange, the quality of the ISF-derived
estimate of blood glucose levels will be determined both by the quality of the
mathematical model on which the calculation of blood glucose levels from
interstitial glucose levels is based and by the accuracy and precision of the
interstitial glucose measurements.
There have been few attempts to assess the transcapillary glucose exchange
properties of the intact peripheral tissues of the rat
(42), dog
(35,
36,
48), and sheep
(13). In these animal studies,
glucose or tracer glucose kinetics simultaneously observed in blood and ISF
have been mathematically treated to yield quantitative information on the
tissue-specific ability to exchange glucose across the capillary walls. To the
best of our knowledge, similar evaluations of transcapillary glucose exchange
in peripheral tissues in humans have not been carried out. Therefore, the
purpose of the present study was to quantify the kinetics of the exchange of
glucose between the plasma and the ISF of human skeletal muscle and adipose
tissue under fasting conditions. To achieve this, we simultaneously sampled
plasma and ISF of muscle and adipose tissue during intravenous infusions of
D-[6,6-2H2]glucose in fasted normal subjects.
ISF sampling from the two tissues was accomplished by employing the open-flow
microperfusion technique (40,
51). The observed tracer
glucose kinetics in plasma and ISF were then analyzed by a capillary-tissue
exchange model to yield kinetic parameters describing transcapillary glucose
exchange in muscle and adipose tissue.
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METHODS
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Subjects. Five male volunteers [age 30.2 ± 2.6 yr, range
2437 yr; body mass index (BMI) 23.4 ± 0.8 kg/m2,
range 21.725.7 kg/m2; means ± SE] participated in
this study. They were all healthy as judged by medical history, physical exam,
and routine laboratory tests and were not taking any medications. Written
informed consent was obtained after the purpose, nature, and potential risks
of the study were explained to the subjects. The experimental protocol was
approved by the ethics committee of the University of Graz.
Open-flow microperfusion and mannitol calibration technique. The
open-flow microperfusion technique has been previously described in detail
(10,
37,
40). Briefly, as described in
these studies, open-flow microperfusion was based on a double-lumen catheter
(18 gauge) with macroscopic perforations (80 holes, each 0.5 mm in diameter).
This catheter was placed in the tissue of interest and then perfused with
artificial extracellular fluid by means of a peristaltic pump (Minipuls 3;
Gilson, Villiers-le-Bel, France). The perfusion fluid entered the probe via
the inlet tubing, moved through the inner cannula to the tip of the probe, and
streamed back in the space between inner cannula and the perforated outer
cannula. The volume of the space between inner cannula and the perforated part
of the outer cannula was 4.8 µl. As the perfusate passed by the
perforations of the outer cannula, exchange occurred between the perfusate and
the surrounding ISF. The medium (now called effluent) then flowed through an
800-mm piece of 250-µm-ID outlet tubing, at the end of which samples were
collected for the subsequent analysis of glucose and other substances. The
total volume from the middle of the perforated region of the outer cannula to
the end of the outlet tubing was 43.5 µl. To achieve an acceptable time
resolution, the microperfusion probes are usually perfused at relatively high
flow rates (e.g., 2 µl/min). However, at these high flow rates, the mixing
between the perfusate and the ISF is not complete. In the present study, we
applied the mannitol calibration technique to determine the extent of the
mixing between perfusate and ISF. In this calibration technique, mannitol is
intravenously infused at a constant rate, and the mannitol concentration is
then measured in the plasma and probe effluents. Mannitol is a sugar with a
composition and molecular weight similar to those of glucose but, unlike
glucose, it is not taken up by muscle and fat cells. Thus steady-state
mannitol concentrations are identical in plasma and ISF of muscle and fat
tissues. Therefore, the extent of the mixing between perfusate and ISF (also
called recovery) can be calculated from the mannitol concentrations in plasma
and probe effluents. The macroscopic perforations of the microperfusion
catheter permit unrestricted exchange of solutes between the perfusion medium
and the surrounding ISF. Because of this property, the microperfusion system
may be qualified to follow rapid solute concentration changes in the ISF
(37,
40).
Intravenous infusions and blood sampling. All subjects were
studied in the supine position after a 12- to 14-h overnight fast. In the
morning at
8 AM, an 18-gauge catheter was placed into a forearm vein for
the infusion of mannitol and tracer glucose. A second catheter was inserted
into a dorsal hand vein on the opposite arm to allow blood withdrawal during
the experiment. The hand with the sampling catheter was placed in a
thermoregulated box and maintained at 55°C to ensure the arterialization
of the venous samples. The sampling cannula was kept patent by the slow
infusion of 0.9% NaCl (Fresenius Kabi Austria, Graz, Austria). A primed (49.5
mg/kg) continuous (0.45 mg · kg-1 · min-1)
infusion of D-mannitol (Fresenius Kabi Austria) was then started at
-120 min and continued for the duration of the study. After a mannitol
equilibration period of 100 min, a basal blood sample (4 ml) was collected at
-10 min. At time 0,
D-[6,6-2H2]glucose (Cambridge Isotope
Laboratories, Woburn, MA) was given as a primed (12 mg/kg) continuous (0.1 mg
· kg-1 · min-1) infusion for 110 min.
During the tracer infusion, 4-ml blood samples were taken at 2.5, 4, 5.5, 7.5,
9.5, 12.5, 17.5, 22.5, 27.5, 35, 45, 60, and 90 min.
ISF sampling. Shortly after insertion of the dorsal hand catheter,
one microperfusion probe was placed in the periumbilical subcutaneous adipose
tissue and another in the tissue of the rectus femoris muscle
(10). Starting at -120 min,
and continuing for the duration of the study, microperfusion probes were
perfused with Ringer solution (Fresenius Kabi Austria) at a constant rate of 2
µl/min. After an equilibration perfusion period of 100 min, effluent
samples were collected continuously according to the following schedule: from
-20 to 0 min, in a 20-min fraction; from 0 to 30 min, in 5-min fractions; from
30 to 50 min, in 10-min fractions; from 50 to 70 min, in a 20-min fraction;
and from 70 to 110 min, in a 40-min fraction. The effluent sampling delay time
introduced by the dead space volume of the probe outlet tubing (21.75 min) was
taken into account when sample collection was begun, and substance
concentrations in the effluent samples were compared with those in the plasma
samples. Separate in vitro experiments have been performed to assess the
extent of dispersion of glucose gradients during the passage of the effluent
through the probe outlet tubing. The results of these experiments suggest
minimal dispersion effects on the shape of the effluent concentration-time
curves.
Sample handling and analysis. Blood samples were placed
immediately in ice and centrifuged at 4°C, and the supernatant was frozen
for assay. Effluent samples were collected in plastic vials (PCR softtube 0.2
ml, Biozyme Diagnostik, Oldendorf, Germany) on ice, capped immediately, and
frozen for assay. During the effluent collection, the vials were covered to
prevent fluid evaporation. To determine the exact sample volume and to monitor
the flow of the perfusate solution, the vials were weighed before and after
collection. The concentrations of glucose and mannitol, as well as the
enrichment of [6,6-2H2]glucose in plasma or effluent
samples, were measured as follows. A 10-µl plasma or effluent sample was
added to 10 µl of internal standard mixture containing
[U-13C]glucose and [1-13C]mannitol (Isotec, Miamisburg,
OH) in known amounts. To this sample, zinc sulfate and barium hydroxide were
added (37). After
centrifugation, the supernatant was passed through a column of mixed-bed anion
and cation ion-exchange resins (Dowex 1-X8200 and Dowex
50W-X8200, Sigma-Aldrich Handels, Vienna, Austria)
(37). The column eluate was
evaporated in a speed vacuum, and the residue was treated with acetic
anhydride and pyridine to convert glucose and mannitol to their acetate
derivatives (6,
7). A Trace GC-MS (Thermo
Finnigan) equipped with an AS 2000 autosampler (Thermo Quest) and a DB-5MS
Capillary GC column (length 15 m, ID 0.25 mm, film thickness 0.25 µm;
J&W Scientific) was used to analyze the glucose and mannitol derivatives.
The GC injector temperature was set at 280°C, and the transfer line
between GC and MS was held at 280°C. The carrier gas was helium. The
column temperature was 80°C for 1 min and was increased by 30°C/min to
310°C. Under chemical ionization (CI), ions with mass-to-charge ratios
(m/z) of 331, 333, and 337 were monitored for glucose, and ions with
m/z ratios of 375 and 376 were monitored for mannitol
(6). The CI gas used was
methane. The peak areas of the monitored ions were used to calculate the
isotope ratios. The variance in the determination of the isotope ratios was
assessed by analyzing replicates of plasma and ISF samples enriched with known
amounts of [6,6-2H2]glucose and of tracers used as
internal standards (i.e., [U-13C]glucose,
[1-13C]mannitol). For example, to assess the variability in the
measurement of the 333/331 isotope ratio (R333/331) in plasma
samples, pooled human plasma with known levels of natural glucose (i.e.,
tracee glucose) and a standard solution of
[6,6-2H2]glucose were used to prepare a series of five
samples ranging in tracer-to-tracee ratio (TTR) from 0 to 12%. Each sample was
aliquoted in five equal volumes. The aliquots were then purified, derivatized,
and analyzed. Over this range of TTR values, the measurement of
R333/331 had a coefficient of variation (CV) equal to 3.2%. A
similar CV in the measurement of R333/331 was obtained for ISF
samples. Furthermore, the R337/331 and R376/375 of the
replicate plasma and ISF samples enriched with known amounts of
[U-13C]glucose and [1-13C]mannitol were determined with
a CV equal to 1.5%. Uncertainties associated with the pipetting are included
in this CV value.
Calculation of glucose and mannitol concentrations in plasma and
effluent samples. Because of the natural occurrence of heavier isotopes
in both tracer and tracee, the isotope ratio cannot generally be taken as a
direct measure of the TTR in a sample. It can be shown
(34,
53) that the general form of
equation defining the relationship between a measured isotope ratio
RL/K and the number of fragment ions arising from a tracee
(Ii) and tracer (Ii*) in a sample is
 | (1) |
where qK and rK are the respective
probabilities of occurrence of tracee and tracer fragment ions with an
m/z of K, and where qL and rL
are the respective probabilities of occurrence of tracee and tracer fragment
ions with an m/z of L. Because all tracers used in our study were 99%
enriched, the contribution of tracer fragment ions at an m/z of K is
negligible (i.e., rK
0). Under this condition, the
following relationship may be deduced from Eq. 1
 | (2) |
Thus, in this case, TTRi (or
Ii*/Ii) is linearly related to
RL/K. Numerical values of the slopes (i.e.,
qK/rL) and intercepts (i.e.,
qL/rL) were obtained by performing
linear regression using data from the GC-MS analysis of the replicate plasma
and ISF samples with known enrichments. There was no deviation from linearity
over the range of TTRi values investigated (r > 0.997).
The measured RL/K values and the determined linear regression lines
(Eq. 2) were then used to derive the TTRi values for the
plasma and effluent samples with unknown enrichments. From the TTRi
values so obtained, contents of unlabeled glucose (G) and mannitol (M) in the
samples were calculated according to the following equations
 | (3) |
 | (4) |
where TTRM is the ratio of [1-13C]mannitol tracer to
mannitol tracee, TTRG is the ratio of [U-13C]glucose
tracer to glucose tracee, and MS* and
GS* are the known quantities of added
[1-13C]mannitol and [U-13C]glucose tracers,
respectively. By carrying out error propagation analysis
(5), it can be shown that the
following relationships between the CVs of the derived tracee concentrations
and those of the measured RL/K values
(CVRL/K) exist
 | (5) |
 | (6) |
where CVM and CVG are the CVs for the determination of
mannitol and glucose concentrations, respectively. The average CVM
for the mannitol data was 2.0% (range 1.73.5%; lower when
R376/375 was higher, see Eq. 5). Because the contribution
of glucose tracee fragment ions at m/z of 337 was negligible (i.e.,
q337
0), the regression line for TTRG
exhibited an intercept at the origin (i.e.,
q337/r337
0). As a result (see
Eq. 6), the CV of the glucose tracee data was close to the
experimentally determined CV for R337/331 (i.e., 1.5%). From the
derived glucose tracee concentrations (G) and the ratios of
[6,6-2H2]glucose tracer to glucose tracee
(TTRD), the content of [6,6-2H2]glucose
(G*) in each plasma or effluent sample was calculated according to
the following equation
 | (7) |
Again, using a propagation of errors analysis, it can be shown that the CV for
the determination of G* (CVG*) can be
calculated as follows
 | (8) |
 | (9) |
The average CVG* values for the determination of
[6,6-2H2]glucose concentrations in plasma and effluent
samples were 5.2% (range 3.911.7%).
Calculation of the ISF glucose concentrations. As already
mentioned, the substrate recovery in the effluent of the microperfusion probe
is not complete at the flow rate used in this study (i.e., the effluent
concentration is lower than the interstitial concentration). However, provided
that the substrate recovery for an effluent sample is exactly known, the ISF
substrate concentrations can be calculated as the substrate concentration in
the effluent divided by the substrate recovery
(40). In the present study,
the substrate recovery for each effluent sample (SR) was estimated as the
ratio of the mannitol concentration in the effluent sample (ME) to
the steady-state concentration of mannitol in the corresponding arterial
plasma sample (MA). Thus, for a certain sampling time point, the
interstitial concentrations of glucose tracee and
[6,6-2H2]glucose tracer were calculated as defined in
Eqs. 10 and 11
 | (10) |
 | (11) |
where GI* and GI are the interstitial
concentrations of [6,6-2H2]glucose tracer and glucose
tracee, respectively. The CVs for the determination of
GI* and GI were calculated from the following
equations
 | (12) |
 | (13) |
CVs of the interstitial [6,6-2H2]glucose tracer data
averaged 6.0% (range 4.612.7%), and the CVs of the interstitial tracee
glucose data averaged 3.2% (range 2.85.2%). Because each probe effluent
sample was collected over a specified time interval (e.g., the effluent sample
in the period from 0 to 5 min), the derived interstitial tracer and tracee
glucose values were considered valid at the midpoint of the interval (e.g., at
2.5 min).
In the calculation of the substrate recovery in the microperfusion probe
effluents (Eqs. 10 and 11), we implicitly assume that
mannitol is not taken up by muscle and fat cells and that, therefore, the
steady-state mannitol concentrations are identical in plasma and ISF of muscle
and adipose tissue. In preliminary experiments in humans, we infused mannitol
intravenously and measured the steady-state mannitol concentration in the ISF
of adipose tissue by combining open-flow microperfusion and the no-net-flux
protocol. According to this protocol
(10,
24), the microperfusion probe
was perfused with different concentrations of mannitol, and the equilibrium
concentration where no net flux of mannitol occurs (i.e., neither
concentration nor dilution of the perfusate mannitol due to the exchange with
ISF mannitol) was determined. We found that this equilibrium concentration of
mannitol in the perfusate was similar to the mannitol concentration in plasma
(Schaupp L, Schaller H, Regittnig W, and Pieber TR; unpublished observation).
Thus, in view of this observation, it seems very likely that the assumption of
identical mannitol concentrations in plasma and ISF of muscle and adipose
tissue is correct.
Analysis of tracer glucose kinetics. In the present investigation,
the analysis of the observed kinetics of tracer glucose is based on a model of
capillary-tissue exchange proposed by Johnson and Wilson
(19). This model assumes that
the interstitial space is well mixed and that the capillaries in the tissue
are evenly distributed and of uniform length. Under these assumptions, Johnson
and Wilson derived equations that describe the intracapillary solute
concentration profiles, as well as the solute concentration in the
interstitial space, as a function of time. Furthermore, these authors showed
that when the interstitial solute concentration is changing slowly compared
with the capillary transit time of an element of blood, the following
relationship may exist (19)
 | (14) |
where P is the capillary permeability, S is the capillary
surface area, Q is the blood flow, and
is a rate constant defined as the
rate of loss of solute from the capillaries per unit concentration difference
between arterial blood and ISF. In the present study, we utilized this
relationship obtained by Johnson and Wilson
(19) to describe the
transcapillary flux of tracer glucose from blood into muscle and adipose
tissue ISF as a function of arterial and interstitial tracer glucose
concentrations. With the additional assumption that there is no back flux of
tracer glucose from the cell, the following differential equation for the
quantities of tracer glucose in the interstitial space can be set up
(52)
 | (15) |
where GA* is the tracer glucose concentration in the
arterial plasma, GI* is the tracer glucose concentration
in the ISF,
0 is the rate constant for uptake of tracer
glucose by the cells, and VI is the distribution volume of glucose
in the ISF space. Dividing the differential equation by VI yields
 | (16) |
where p1 =
/VI, and p2
= (
/VI =
0/VI). Both parameters
are uniquely identifiable from plasma and ISF tracer data. A graphic
visualization of the mathematical model defined with Eq. 15 is given
in Fig. 1. To characterize the
specific properties of the blood-tissue exchange of glucose in the examined
tissues, the steady-state ratio of ISF glucose to plasma glucose (r)
and the 95% equilibrium time (T95%) were calculated from
p1 and p2 as follows
 | (17a) |
 | (17b) |
The time constant T95% is defined as the time required for the ISF
glucose to attain 95% of the steady-state value after a concentration step
challenge in the plasma compartment. Thus the value of T95%
provides a measure of the ISF glucose equilibration delay, which is determined
by the fractional rate of glucose exchange (
/VI) and the
fractional rate of glucose uptake (
0/VI).
Numerical methods. To perform the kinetic analysis, the observed
plasma tracer profiles (GA*) were used as an input to
Eq. 16, and the parameters (p1,
p2) were identified by fitting the output
(GI*) to the observed ISF tracer data. When plasma
tracer data were used as an input, the plasma concentrations between sample
times were determined by linear interpolation. The numerical values of unknown
parameters were obtained by weighted nonlinear least squares by use of a
Levenberg-Marquadt algorithm
(37) with inverse variance
weights. Monte Carlo analysis was performed to determine the precision of the
parameter estimates (37). The
measurement errors were assumed to be independent, Gaussian of zero mean, and
with the experimentally determined variances. The statistical significance of
differences was calculated using two-tailed, paired Student's t-test
analysis. A P value of <0.05 was considered to indicate
statistical significance. Data are reported as means ± SE. Steady-state
values were calculated by averaging data from the final hour of the
experiment. MATLAB software packages (The MathWorks, Natick, MA) were used for
all of the analyses and statistics
(37).
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RESULTS
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Mannitol concentrations and substrate recovery. In the five
subjects studied, the primed-continuous infusion of mannitol resulted in a
constant plasma mannitol concentration of 1.62 ± 0.12 mmol/l
(Fig. 2A). In
comparison, the mannitol concentrations in the effluents of the muscle and
adipose tissue probes attained a steady-state level of 0.35 ± 0.07 and
0.27 ± 0.05 mmol/l, respectively
(Fig. 2A). By taking
these mannitol values, the average substrate recovery in the muscle and
adipose tissue probe effluents was calculated to be 21.1 ± 2.9 and 17.4
± 3.4%, respectively. There was no statistical difference in substrate
recovery between the two probe effluents (P > 0.32). The
relatively constant mannitol concentrations in plasma and effluent samples
indicate that steady-state microperfusion conditions were maintained during
the experiments.
Total glucose concentrations. During the final hour of the study,
the average total glucose concentrations (i.e., tracer + tracee
concentrations) in the muscle and adipose tissue probe effluents were 0.670
± 0.065 and 0.609 ± 0.121 mmol/l, respectively
(Fig. 2B). By taking
the observed effluent glucose and substrate recovery values (Eqs. 10
and 11), the mean total glucose concentrations in muscle and adipose
tissue ISF can be calculated to be 3.32 ± 0.46 and 3.51 ± 0.17
mmol/l, respectively. There was no statistical difference in concentrations
between muscle and adipose tissue ISF. However, the total glucose levels
estimated for ISF of muscle and adipose tissue were significantly lower than
those measured in arterialized plasma (5.56 ± 0.13 mmol/l, P
< 0.01). Thus, in these subjects, a significant glucose gradient between
plasma and ISF of muscle and adipose tissue was observed in the fasting state
(Fig. 2B).
Glucose TTR. Figures
3A and
4A show the time
courses of the glucose TTR in plasma and ISF of muscle and adipose tissue. As
can be seen, the glucose TTR measured in plasma increased to a peak of 0.118
± 0.021 at 2.5 min after start of infusion, fell for
45 min to a
level of 0.055 ± 0.004, and remained at this level until the end of the
experiment. In the muscle ISF, the glucose TTR increased gradually during the
first
20 min of infusion and then reached a steady-state level comparable
to that seen in plasma (Fig.
3A). A similar pattern of change in the glucose TTR was
observed in the ISF of adipose tissue (Fig.
4A). The marked dynamic differences between the glucose
TTR in plasma and those in the ISF of the two tissues suggest that, at
20
min after the start of the tracer glucose infusion, the plasma and the
interstitial tracer glucose pools attained their exchange equilibrium.
Tracer glucose concentrations and kinetic analysis. As can be seen
in Figs. 3B and
4B, the tracer glucose
concentration in plasma rose to a peak of 0.67 ± 0.13 mmol/l by 2.5 min
after start of infusion, declined for
45 min, and then remained at a
steady-state level of 0.29 ± 0.03 mmol/l. In comparison, interstitial
tracer glucose concentrations in both muscle and adipose tissue gradually
increased during the first
20 min of tracer infusion. After this period,
tracer glucose concentrations in muscle and adipose tissue ISF paralleled
those seen in plasma. During the final hour of the study, the tracer glucose
concentrations in muscle and adipose tissue ISF averaged 0.19 ± 0.04
and 0.20 ± 0.02 mmol/l, respectively. In each subject, tracer glucose
profiles in plasma and ISF were used to identify the parameters of the
capillary-tissue exchange model (Fig.
1). Tables 1 and
2 list the individual
parameters derived from the observed exchange kinetics in muscle and adipose
tissue, respectively. Parameter CVs as determined by Monte Carlo analysis
ranged from 9 to 16%. Thus parameters of the proposed model were estimated
with a relatively high degree of precision. Average model fits for tracer
glucose kinetics observed in muscle and adipose tissue ISF are shown in Figs.
3B and
4B, respectively. As
can be seen, interstitial tracer glucose kinetics in both tissues were well
described by the capillary-tissue exchange model. The value of the model
parameter p2 derived from adipose tissue data was similar
to the value of p2 derived from muscle data (0.120
± 0.015 vs. 0.110 ± 0.012 min-1, P >
0.57). When the values of p2 are used, the 95% equilibrium
times (T95%) for skeletal muscle and adipose tissue were calculated
to be 28.6 ± 3.2 and 26.8 ± 3.6 min, respectively. As was the
case with parameter p2, the value of
p1 derived from muscle data did not differ from the value
identified from adipose tissue data (0.063 ± 0.007 vs. 0.081 ±
0.012 min-1; P > 0.31). By taking the values of
p1 and p2, the steady-state ratios of
ISF to plasma glucose can be calculated to be 0.58 ± 0.06 and 0.66
± 0.03 for skeletal muscle and adipose tissue, respectively. There was
no significant difference in the ratios between the two tissue beds
(P > 0.38).
 |
DISCUSSION
|
---|
The objective of this investigation was to assess transcapillary glucose
exchange efficiency in human skeletal muscle and adipose tissue under basal
conditions. For this reason, we frequently sampled plasma as well as muscle
and adipose tissue ISF during intravenous
[6,6-2H2]glucose infusion in fasting, nonobese humans.
The ISF sampling from the two tissues was accomplished by applying the
open-flow microperfusion technique
(40). To achieve a relatively
high time resolution with this technique, a perfusion flow rate of 2 µl/min
was employed. At this high flow rate, the mixing between the perfusate and the
ISF surrounding the probe is not complete. Therefore, to determine the extent
of perfusate-ISF mixing in the probe effluents (i.e., recovery), a constant
intravenous infusion of the extracellular marker mannitol was initiated 120
min before the beginning of tracer infusion and was continued throughout the
experiment. In this way, a constant plasma mannitol level was established
during the plasma and ISF sampling period
(Fig. 2A). Because
steady-state concentrations of mannitol are similar in plasma and ISF of
muscle and adipose tissue (see METHODS), the observed
effluent/plasma ratios of mannitol were taken as estimates of substrate
recovery in the collected probe effluents. From the substrate recovery and
glucose values in the probe effluents, we were then able to determine the
interstitial glucose concentrations in muscle and adipose tissue. We found
that, in the fasting state, the glucose levels in the ISF of both adipose
tissue and skeletal muscle are
60% of arterialized plasma levels
(Fig. 2B). This result
suggests that a significant glucose concentration gradient between plasma and
ISF of muscle and adipose tissue exists in the fasting state.
In a recent microperfusion study, by employing different calibration
techniques, we found similar glucose concentrations in adipose tissue ISF in
normal subjects under fasting conditions
(40). Furthermore, some other
investigators have measured interstitial glucose levels in human adipose
tissue using microdialysis
(26,
29,
32,
41), ultrafiltration
(41), and/or an equilibration
technique (41) and have also
reported glucose levels in the ISF to be significantly lower than those in
plasma, with levels ranging from 50 to 75% of arterialized plasma values.
However, contrary to these results, a number of studies using microdialysis in
normal subjects have estimated the basal interstitial glucose concentrations
in adipose tissue to be 85100% of arterialized plasma levels
(4,
18,
24,
28,
39,
47,
50). Concerning the human
skeletal muscle, previous studies have employed the microdialysis technique to
measure glucose levels in the muscle ISF and have found fasting interstitial
glucose concentrations to be equal to
(25,
26,
30,
31) or higher [85100%
of arterialized plasma values
(7,
15,
28,
32,
39)] than those measured in
the present microperfusion study. Although spatial heterogeneity in the
interstitial glucose levels within a studied tissue bed (e.g., due to local
variations in blood flow) cannot be excluded, it is reasonable to suspect that
technical and procedural details can account for the different findings. For
example, Hamrin et al. (14)
have reported that the hydrostatic pressure applied to the perfusate of the
microdialysis catheter influences the fluid and substance transport across the
microdialysis membrane. Hydrostatic pressure changes induced by varying the
vertical position of the orifice of the microdialysis outlet tubing (i.e.,
below or above the inlet of the microdialysis catheter) resulted in
significant changes in the dialysate substrate concentrations, as well as in
substantial variations in the degree of perfusate losses (due to net perfusate
flux across the membrane into the tissue). Thus the divergent findings of
previous microdialysis studies may be partially attributable to different
hydrostatic pressures applied to the perfusate of the microdialysis probes.
Counteracting the hydrostatic pressure by adding osmotically active substances
to the perfusate (e.g., dextran-70) may abolish this source of error
(14).
In the present study, the initiation of the intravenous tracer glucose
infusion in the primed-continuous format produced a rapid rise in the plasma
tracer glucose concentration (Figs.
3B and
4B). After attaining
peak values during the first 2.5 min of tracer infusion, plasma tracer
concentration fell, first rapidly and then more slowly until, after
45
min, the plasma level remained constant for the remainder of the experiments.
In contrast to the rapid rise and fall with an early, sharp peak seen in the
plasma tracer levels, the interstitial tracer concentration in both muscle and
adipose tissue rose more slowly to a smooth peak at
20 min after the
tracer infusion was begun. Subsequently, the tracer glucose concentrations in
ISF of muscle and adipose tissue paralleled those seen in plasma (Figs.
3B and
4B). Similar dynamic
differences in the time course of the TTR between the plasma and ISF of muscle
and adipose tissue were observed during the first
20 min of the
experiments (Figs. 3A
and 4A). These dynamic
differentials seen between the plasma and interstitial tracer levels, as well
as the plasma and interstitial TTRs, strongly suggest that, after
administration of a primed-continuous tracer glucose infusion, the process of
tracer equilibration between plasma and ISF of adipose tissue and skeletal
muscle takes
20 min.
Previous studies in humans have shown that the disappearance of labeled
glucose from plasma can be described by the sum of two or more exponential
components (8,
11,
16,
17,
37,
43). The different exponential
components of the tracer disappearance curve have then usually been
interpreted as indicating diffusion from plasma into different anatomic fluid
compartments, such as interstitial and intracellular fluids of various organs
and tissues (17). For example,
Ferrannini et al. (11) and
Cobelli et al. (8) have studied
in detail the plasma dynamics of injected labeled glucose in normal subjects,
both in the fasting state and in the high-insulin euglycemic state. The
authors found that, under both experimental conditions, the plasma
disappearance curves of labeled glucose were best described by the sum of
three exponentials, thus indicating the presence of two separate compartments
exchanging glucose with the plasma compartment at two different rates. The
initial rapid exponential decline observed during the first
2 min after
tracer injection was attributed to fast tracer diffusion from plasma into
extravascular fluid spaces of liver, spleen, and endocrines, whereas the decay
of the slower component observed during the first 2030 min was
interpreted as indicating slow diffusion into the extravascular fluid spaces
of insulin-sensitive tissues, such as the interstitial space of skeletal
muscle and adipose tissue (8,
11). These interpretations
given by the authors are supported by the well-known facts that both the
capillary permeability and blood supply are considerably lower in skeletal
muscle and adipose tissue than in splanchnic organs
(9). Because of the technical
difficulties in obtaining observational access to the ISF compartment of human
skeletal muscle and adipose tissue, this physiological meaning assigned to the
slower exponential component of the plasma tracer disappearance curve has not
been previously verified in vivo. However, in the present study, by combining
open-flow microperfusion with the mannitol calibration technique and a
sensitive glucose-measuring method (GC-MS), we were able to assess
interstitial glucose kinetics in muscle and adipose tissue with an acceptable
time resolution. Our observations of marked dynamic differentials between
plasma and interstitial tracer glucose in muscle and adipose tissue during the
first
20 min of intravenous tracer infusions (Figs.
3B and
4B) thus provide
direct experimental evidence that the interstitial glucose pool of muscle and
adipose tissue is indeed part of the postulated compartment that is in slow
equilibrium with plasma glucose.
The constancy of the plasma tracee glucose concentration during the
experiments (data not shown) and the similarity of the pattern of change in
the tracer glucose concentration and TTR (Figs.
3 and
4) indicate that the endogenous
glucose system was not perturbed from its steady state by the tracer infusion.
This condition allowed us to apply a published capillary-tissue exchange model
for the analysis of tracer glucose kinetics
(Fig. 1). This model was first
proposed by Johnson and Wilson
(19) to describe the
transcapillary exchange kinetics of nonmetabolizable tracers and was later
modified by Watson (52) to
include the effects of cell uptake for metabolizable tracers. Mathematical
models of more complex form have been proposed for the description of
capillary-tissue exchange. However, on the basis of fewer simplifying
assumptions, these models contain many more adjustable parameters than can
possibly be determined experimentally in humans (see Ref.
3 for review). The
capillary-tissue exchange model used in the present study represents the
interstitial space in terms of a well-mixed compartment
(Fig. 1) and, thus, implicitly
assumes that glucose diffusion in the ISF is an instantaneous process. The
conception of the ISF of muscle and adipose tissue as single well-mixed
compartments is a considerable simplification, but it may be justified by the
fact that, in muscle and adipose tissue, the longest intercapillary diffusion
distance is 3050 µm
(9), and that glucose diffusion
over such a distance is very rapid [within a few seconds
(9,
21)]. Also implicit in the
model is the assumption that the capillary transit time of an element of blood
is short compared with time constants of the tracer equilibration between
blood and ISF (19,
52). This assumption seems to
be appropriate for the mathematical treatment of the transcapillary glucose
exchange process in muscle and adipose tissue, because the capillary transit
time is on the order of 12 s
(9), and the observed 95%
equilibrium time of the interstitial glucose is on the order of 30 min in
these tissues (Tables 1 and
2; Figs.
3 and
4). The model further assumes
that tracer is phosphorylated as rapidly as it is translocated into the cell,
so that back flux of tracer from the cell is negligible. This latter
assumption appears to be justified in our experimental situation, because
myocytes and adipocytes may have very little or no free intracellular glucose
under basal euglycemic conditions
(6,
7,
12,
20).
To perform the kinetic analysis, the interstitial tracer glucose
concentrations were fitted with the capillary-tissue exchange model by using
the plasma tracer profile as an input. As can be seen in Figs.
3B and
4B, the interstitial
tracer glucose kinetics in both muscle and adipose tissue were well described
by the model, and, as indicated by the low CV values, the model parameters
were estimated with a relatively high degree of precision (Tables
1 and
2). To obtain a measure of the
transcapillary glucose exchange efficiency in skeletal muscle and adipose
tissue, the 95% equilibrium time (T95%) was calculated from the
identified model parameters. We found that the value of T95% is
28.6 ± 3.2 min for skeletal muscle and 26.8 ± 3.6 min for
adipose tissue. There was no significant difference in the T95%
value between the two tissues (P = 0.60). The mean values of
T95% estimated in the present study are lower than those found by
Steil et al. (48) and Rebrin
and Steil (35), who analyzed
the hindlimb lymph kinetics of labeled glucose in the dog under basal and
hyperinsulinemic conditions. These authors observed a T95% value of
36.9 ± 7.2 min in the basal state and 40.4 ± 6.9 min in the
hyperinsulinemic state [T95% is related to the reported
T63% as T95% = -ln(0.05) T63%
(35)]. Because hindlimb lymph
fluid is primarily derived from the ISF of skeletal muscle
(48), the T95%
values observed in this previous lymph study may therefore largely reflect the
transcapillary glucose exchange efficiency of skeletal muscle in dogs. In a
more recent study, Rebrin et al.
(36) have measured
interstitial glucose kinetics by means of a glucose sensor implanted in the
subcutaneous tissue of the dog and have observed T95% values (range
1037 min) that are on the average similar to those derived in our
study. Furthermore, in a previous review article, Crone and Levitt
(9) provided a summary of
permeability-surface area product (PS) values that had been obtained
by various investigators using different measurement methods. From the
PS values, Crone and Levitt calculated T95% values for
skeletal muscle and adipose tissue. These T95% values (range
930 min) are on the average lower than those obtained in the present
and previous (36,
48) in vivo studies. However,
as pointed out by Crone and Levitt, the values of T95% were
calculated with the assumption of permeability-limited exchange (i.e., no
influence of the blood flow), and thus may not reflect the actual in vivo
situation, where both the capillary permeability and the prevailing blood flow
determine the transcapillary glucose exchange rates.
In summary, our results obtained using open-flow microperfusion demonstrate
that, in the fasting state, the glucose levels in the ISF of human skeletal
muscle and adipose tissue are
60% of arterialized plasma levels.
Furthermore, the mathematical analysis of the interstitial tracer glucose
kinetics measured in muscle and adipose tissue during intravenous tracer
infusions indicates that the 95% equilibrium time for the two tissues is
28 min. Thus the observed ISF-to-plasma glucose gradients and derived 95%
equilibrium times provide strong evidence that transcapillary exchange of
glucose is limited in human skeletal muscle and adipose tissue. The
interstitial glucose pool of muscle and adipose tissue may therefore be
considered as part of a compartment that is in relatively slow equilibrium
with plasma glucose.
 |
DISCLOSURES
|
---|
This study was supported by the Austrian Science Fund, Project P16228
[GenBank]
-B05,
and the European Community, Project ADICOL-IST-199914027.
This work was presented at the 62nd Scientific Sessions of the American
Diabetes Association, June 1418, 2002, San Francisco, California.
 |
ACKNOWLEDGMENTS
|
---|
We thank C. Schlack and Dr. W. Windischhofer for their help with the
measurement of the stable isotope, and Dr. G. A. Brunner for assistance in the
performance of the experiments.
 |
FOOTNOTES
|
---|
Address for reprint requests and other correspondence: W. Regittnig, Dept. of
Biophysics, Institute of Biomedical Engineering, Graz Univ. of Technology,
Krenngasse 37, A-8010 Graz, Austria (E-mail:
werner.regittnig{at}healthgate.at).
Submitted 9 August 2002
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. Section 1734
solely to indicate this fact.
 |
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