Departments of 2 Biochemistry, 3 Medicine, and 1 Nutrition, Case Western Reserve University School of Medicine, Cleveland, Ohio 44106
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The contribution of gluconeogenesis to glucose production can be measured by enriching body water with 2H2O to ~0.5% 2H and determining the ratio of 2H that is bound to carbon-5 vs. carbon-2 of blood glucose. This labeling ratio can be measured using gas chromatography-mass spectrometry after the corresponding glucose carbons are converted to formaldehyde and then to hexamethylenetetramine (HMT). We present a technique for integrating ion chromatograms that allows one to use only 0.05% 2H in body water (i.e., 10 times less than the current dose). This technique takes advantage of the difference in gas chromatographic retention times of naturally labeled HMT and [2H]HMT. We discuss the advantage(s) of using a low dose of 2H2O to quantify the contribution of gluconeogenesis.
diabetes; stable isotopes; gas chromatography-mass spectrometry
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
INVESTIGATORS HAVE PROPOSED numerous isotope tracer methods (i.e., precursor-to-product labeling techniques) for quantifying the contribution of gluconeogenesis to glucose production. Unfortunately, the use of carbon-labeled tracers is affected by isotope exchange and/or dilution and by metabolic zonation of the liver (9, 14). Consequently, data from studies that use carbon-labeled tracers are often questionable. The 2H2O method is not subject to such artifacts, and it has been widely used to measure gluconeogenesis (2-8, 12). The principle of the 2H2O method is straightforward. Briefly, after an oral bolus of 2H2O is administered, the contribution of gluconeogenesis to glucose production is determined by measuring the ratio of 2H that is bound to carbon-5 vs. carbon-2 of glucose (2-8, 12).
The current protocol(s) for using 2H2O requires that body water be enriched to ~0.5% 2H. This level of enrichment in body water is necessary to achieve detectable levels of 2H in blood glucose when standard gas chromatography-mass spectrometry techniques are used. The ability to use a lower dose of 2H2O (e.g., 0.05% 2H in body water) could offer certain advantages. First, some patients experience vertigo and/or nausea immediately after drinking 2H2O. Although this represents a minor fraction of subjects given 2H2O (e.g., ~10%; Ref. 5) and seems to depend on how rapidly 2H2O is administered (5, 6), use of a "low dose" of 2H2O should eliminate those side effects. Second, administering less isotope will reduce the cost of performing experiments. Third, because the half-life of body water is ~2 wk, one should allow ~2 mo before repeating measurements of gluconeogenesis in the same subject. This is necessary because blood glucose and liver glycogen remain labeled in the presence of labeled body water (13). If one could measure the contribution of gluconeogenesis on one day by administering a low dose of 2H2O, then it is conceivable that one could measure gluconeogenesis on a later day (e.g., the following day) by administering a "standard dose" of 2H2O. This two-step approach would nullify any contribution of labeled glucose or glycogen; i.e., the higher dose of 2H2O given on the second day would cancel the effect(s) of giving a low dose of 2H2O on the first day. If correct, investigators could, for example, determine the efficacy of a therapy on gluconeogenesis in a relatively acute setting.
We hypothesized that measurements of 2H enrichment could be enhanced by taking advantage of the fact that, during gas chromatography, 2H-labeled molecules typically fractionate from their respective unlabeled forms. If true, gluconeogenesis could perhaps be measured using a low dose of 2H2O. We tested our hypothesis and observed that, in fact, biasing the ion chromatogram integration routine (i.e., measuring the isotope ratio across a limited region of the chromatographic peaks) enhances the detection of 2H. Biological experiments were then performed to determine whether our chromatographic integration method could be used to extend the application of 2H2O. We show that our approach to integrating ion chromatograms allows one to measure gluconeogenesis by using only 0.05% 2H in body water.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Chemicals
Unless specified, all chemicals and reagents were purchased from Sigma-Aldrich. [1-2H]glucose (98 atom percent) and [1-13C]glucose (99 atom percent) were purchased from Cambridge Isotopes (Andover, MA). 2H2O (99 atom percent) was purchased from Isotec (Miamisburg, OH). Gas chromatography-mass spectrometry supplies were purchased from Alltech (Deerfield, IL).Standards of [2H]hexamethylenetetramine (HMT) and [13C]HMT were generated from [1-2H]- and [1-13C]glucose, respectively (3, 16). Briefly, labeled glucose is reduced with NaBH4 to sorbitol. Oxidation with periodic acid generates formaldehyde from carbons 1 and 6 of sorbitol; thus the enrichment of the formaldehyde is one-half that of the original glucose. After distillation, the formaldehyde is reacted with NH4OH to yield HMT.
Biological Experiment
The contribution of gluconeogenesis to glucose production was measured in male Sprague-Dawley rats (Harlan Laboratories, ~250 g). On day 1 (10 AM), rats were injected (3.25 ml/kg ip) with saline or 10% 2H2O-saline (assuming that body water accounts for 65% of body weight, body water should be enriched to ~0.05% 2H). Rats were returned to their cages and allowed free access to food and water. On day 2 (2 AM), food was removed. At 10 AM, rats were injected (3.25 ml/kg ip) with 10 or 99% 2H2O-saline to achieve enrichments of ~0.05 or ~0.5% 2H in body water, respectively. At 3 PM, rats were killed, and total blood was collected.Plasma glucose concentration was determined using a YSI 2300 Stat Plus (Yellow Springs Instrument, Yellow Springs, OH). The labeling of 2H bound to carbons 2 and 5 of blood glucose was determined (3, 16). Briefly, carbons 2 and 5 are isolated as formaldehyde and reacted with NH4OH to yield HMT. The contribution of gluconeogenesis to glucose production was calculated from the ratio of [2H]carbon-5glucose to [2H]carbon-2glucose.
Analyses of HMT
Data acquisition. Gas chromatography-mass spectrometry analyses of HMT were performed using an Agilent 5973N-MSD equipped with an Agilent 6890 GC system. A DB17-MS capillary column (30 m × 0.25 mm × 0.25 µm) was used in all analyses. The temperature program was as follows: 100°C initial, hold for 2 min, increase by 20°C/min to 220°C, and hold for 4 min. The split ratio was 15:1 with a helium flow of 1 ml/min. HMT elutes at 6.1 min. The mass spectrometer was operated in the electron impact mode.
Unless specified, the mass spectrometer was set to monitor two ions [i.e., mass-to-charge ratio (m/z) 140 and m/z 141], at dwell times of 10 ms each. This permits the collection of 84 data points across each ion chromatogram compared with collection of 22 data points across each chromatogram when the dwell time is set to 100 ms/ion [Note that the number of data points is also dependent on other factors, including the absolute quantity of HMT injected and the threshold used to define the signal-to-noise ratio (signal/noise). For purposes of a simplified discussion, the same test sample and threshold settings were used to evaluate the impact of dwell time.].Manual integration. Before the integration was performed, all chromatograms were manually inspected to determine whether the abundance of HMT was sufficient and similar. The peak heights of m/z 140 and m/z 141 are typically in the range of 1,000,000 and 80,000, respectively; the baseline trace of each ion is usually in the range of 100. The raw data (scan number and ion abundance of m/z 140 and m/z 141) were then downloaded to a conventional spreadsheet. The following rationale was used to define start-stop points in the integration routine, and this process was performed in every sample. First, the m/z 140 signal at each scan number was converted to a fraction of the largest m/z 140 signal. Second, the start point of the integration was defined as the scan number at which the m/z 140 signal is 2% of the maximum. Starting at this scan number is necessary to ensure that the signal for m/z 141 is at least ~10 times its baseline. In some analyses, a scan equal to 2% of the maximum could not be identified; in those cases, the start point was defined as the point closest to 2%. The stop point was varied to set increasing degrees of signal amplification. Again, in some instances a stop point at a specific fraction of the largest signal could not be identified. In those cases the stop point was set at the point closest to that where the integration was to be terminated.
Analyses of body water labeling. The 2H enrichment of water was assayed by exchange with acetone, as described by Yang et al. (18), with minor modifications. First, it was not necessary to decrease the ionization energy below 70 eV to obtain a correct basal (M + 1)/(M + 0) signal ratio in acetone. Presumably, the ionization effect(s) reported by Yang et al. do not occur on our instrument. Second, assays were performed using 40 µl of sample or standard, 2 µl of 10 N NaOH, and 4 µl of a 5% (vol/vol) solution of acetone in acetonitrile.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
HMT is a polymer made from six formaldehyde and four ammonia
molecules. Generating HMT from a mixture of labeled and unlabeled formaldehyde yields molecules ranging in molecular weight from 140 to
146. Figure 1 shows the ion chromatograms
of HMT made from naturally labeled, 50% [2H]-, or 50%
[13C]formaldehyde (data were acquired using selected ion
monitoring of m/z 140-146, 10 ms dwell time). In Fig.
1A, the abundance of HMT at m/z 142-146 is
too low to appear on the trace. The elution profile of those mass
isotopomers follows the same pattern (retention time) as HMT of
m/z 140 and 141. Figure 1, B and C,
demonstrates that fractionation of HMT during capillary gas
chromatography is dependent on the type of isotope and not on the
molecular weight; as the molecular weight of HMT made from
[2H]formaldehyde increases, the scan number (retention
time) decreases (Fig. 1B). This does not occur when HMT is
made from [13C]formaldehyde (Fig. 1C).
|
Figure 2 shows that isotope fractionation
is detectable at low 2H enrichments. For example, HMT made
from naturally labeled formaldehyde m/z 141 coelutes
with m/z 140 (Fig. 2A). However, when HMT is made
from 0.05% [2H]formaldehyde, the overlay of the ion
chromatograms is distorted; m/z 141 elutes before
m/z 140 (Fig. 2B).
|
To determine whether the integration method could be used to affect the
response factor, standards of [2H]HMT were analyzed, and
regression plots of the measured-to-expected isotope ratios were made.
Figure 3 shows a representative ion chromatogram with points indicating the cutoffs used for quantitation of enrichment (Fig. 3A). In theory, the labeling of HMT is
about six times that of the formaldehyde. Integrating the entire areas of the peaks of m/z 140 and m/z 141 yields a
slope that is close to the theoretical (Fig. 3B, I).
However, as the chromatographic peaks are cut into smaller units, the
slope of the measured-to-expected isotope ratio increases (Fig.
3B, I-VI). Biasing the integration procedure increases
the detection of [2H]HMT ~3.5 times (Fig. 3B,
I vs. VI).
|
Table 1 shows the coefficient of
variation for measurements displayed in Fig. 3. In all cases, the data
were highly reproducible, and all values were <1%. These data show
that, despite the use of fewer data to measure the isotope labeling
ratio, there is no increase in the error.
|
The contribution of gluconeogenesis to glucose production was measured
in rats that were given 2H2O. Table
2 shows that the enrichment of body water
was ~10 times greater in group 1 than in group
2. In both cases, the absolute enrichment was close to the
expected value based on the administered dose of
2H2O. The enrichment of body water in
group 3 was approximately equal to the sum of the
enrichments of body water observed in groups 1 and
2, as expected. Gluconeogenesis was calculated by comparing
the enrichment of the 2H that is bound to
C-5glucose with that which is bound to
C-2glucose. The 2H enrichment of HMT generated
from C-5glucose and C-2glucose was determined
by integrating only the first 10% of the peak area (this is equivalent
to Fig. 3B, IV). The enrichment was computed by
comparing the data against known standards that were also integrated across the same areas (i.e., the first 10% of the peak). In all cases,
the 2H-labeling on C-2glucose almost completely
equilibrated with that in body water (Table 2). The contribution of
gluconeogenesis was similar in all groups (Table 2).
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The contribution of gluconeogenesis to glucose production can be measured using 2H2O (2-8, 12). The 2H2O method requires that one measure the ratio of 2H that is bound to carbons 5 and 2 of plasma glucose. This is done by isolating carbons 5 and 2 of glucose as formaldehyde and then reacting with ammonia to generate HMT. The 2H enrichment of HMT is then typically calculated after the entire chromatographic peak area of the labeled and unlabeled HMT molecules is measured.
As is often the case for 2H-labeled compounds (11, 17), [2H]HMT elutes ahead of naturally labeled HMT during gas chromatography (Figs. 1 and 2). We hypothesized that focusing the integration on the leading edge of the chromatographic peaks would enhance measurements of 2H enrichment; i.e., biasing the ion chromatogram integration routine would increase the response factor when the measured-to-expected isotope ratio is calculated.
To examine the effect of the integration routine on the detection of 2H, HMT standards of known enrichment were analyzed. We found that limiting the integration to the leading edge of the gas chromatography peaks increases the detection of 2H (Fig. 3) without affecting the reproducibility of the measurements (Table 1). Two points should be considered. First, most of the 21-fold amplification in the enrichment of formaldehyde is due to the polymerization reaction. For example, Fig. 3 shows that our integration procedure amplified the labeling up to 3.5-fold (Fig. 3B, I vs. VI). Second, most likely, highly reproducible measurements of the ratio m/z 141/(140 + 141) were obtained (Table 1) because of the relatively large signal/noise in our analyses (see below).
On the basis of the data in Fig. 3 and Table 1, we hypothesized that one could quantify the contribution of gluconeogenesis with only ~0.05% 2H in body water (i.e., ~10 times less than the dose that is currently used). Biological experiments were performed to address two questions. First, is it possible to measure the contribution of gluconeogenesis if body water is enriched to ~0.05% 2H? Second, is it possible to measure gluconeogenesis on consecutive days by administering the low dose of 2H2O on one day and administering the standard dose of 2H2O on a later day?
Rats in group 1 were given the standard dose of 2H2O (i.e., ~0.5% 2H) and thus served as a control for groups 2 (low dose) and 3 (low dose followed by standard dose). The contribution of gluconeogenesis to glucose production was similar in all groups (Table 2). These data demonstrate that a low dose of 2H2O and a two-step approach can be used to measure the contribution of gluconeogenesis to glucose production. Finding that gluconeogenesis contributes ~60% to glucose production ~13 h after feeding is reasonable. For example, at this time, the concentration of hepatic glycogen is approximately half-maximal (1). As shown by Rothman et al. (15), when hepatic glycogen levels are approximately half-maximal, gluconeogenesis contributes ~50-60% to glucose production.
Using our selective ion chromatogram integration routine raises several points. First, the maximal signal (i.e., the maximal peak height) for m/z 140 was ~10,000 times its baseline. This ensured that, when the integration at 2% of the maximum signal of m/z 140 was started, the signal for m/z 141 was ~10 times its baseline. We found that, when integrations are started before the m/z 140 signal reaches more than 2% of the peak maximum (i.e., when the m/z 141 signal is less than 10 times its baseline), the computed data become erratic, since the signal approaches the noise level. Establishing conditions where the m/z 140 signal is 10,000 times its baseline did not present a problem, since the preparation of HMT (3, 16) generally yields more than enough material. The method will tolerate more, or less, signal/noise. In cases where the signal/noise is >10,000, and with the assumption that the peak shape is not distorted, no major precautions are required. In cases where the signal/noise is <10,000 (e.g., 7,500), one may need to adjust the starting point. For example, integrations may need to be started at ~5% of the maximal signal of m/z 140. However, if integrations are started at points much greater than 2% of the peak maximum, one loses some of the benefit obtained via the chromatographic resolving power. This will depend on the degree of isotope fractionation and the elution profile of the labeled and unlabeled molecules. Of course, if the signal/noise drops too low, samples may need to be reanalyzed.
Second, the degree of isotope fractionation affects the ability to amplify the 2H enrichment. Future experiments are aimed at testing variables that affect isotope fractionation (e.g., column length, linear velocity, etc.).
Last, Matthews and Hayes (10) recognized that quantifying ion beams generated using gas chromatography as an inlet source potentially introduces error(s), since the ion current varies with analyte elution. They noted that the number of scans required to define an ion chromatogram(s), and therefore measure an isotope ratio, requires consideration. They showed that errors in measuring isotope ratios can be minimized by using short dwell times. In our studies, all samples were analyzed using a 10-ms dwell time per ion. We observed that integrating between 2 and 10-25% of the peak maximum (Fig. 3B, III and IV) yields sufficient amplification while still including numerous data points (~10 and 22 points, respectively). Surprisingly, we also obtained reproducible measurements of isotope ratios by integrating the ion current over as few as four data points (Fig. 3B, VI). Although that observation is intriguing, we do not necessarily advocate using an extremely limited amount of data (again, it should be emphasized that the number of data points is dependent on the dwell time and the total quantity of material that is injected). Investigators should evaluate the limits of detection and the reproducibility required for particular analyses.
In their report, Matthews and Hayes (10) also discussed the fact that the degree of isotope fractionation affects the number of data points required to accurately measure an isotope ratio. They noted that, as isotope fractionation increases, a greater number of measurements is required to correctly define the area(s) of a chromatographic peak(s) and accurately quantify an isotope ratio. Those considerations are especially relevant when quantifying high enrichments or when using multiply labeled tracers (e.g., [2H5]glycerol), i.e., conditions where there are clear shifts in the baseline elution of the labeled and unlabeled compounds. Using singly labeled 2H tracers at very low enrichment, there is virtually no detectable shift in the initial elution point for the isotope (Fig. 2). Because the presence of small amounts of 2H affect mainly the leading edge of the chromatographic peak (Fig. 2), it does not appear that extra precautions are necessary for defining the integration start point when measuring low enrichments. We should emphasize that a difference between our approach and the concerns raised by Matthews and Hayes centers on the use of calibration standards; Matthews and Hayes' approach is ideal for measuring enrichment in the absence of calibration standards, whereas our method has an absolute requirement on the use of calibration standards since the measured-to-expected isotope ratio is sensitive to the number of scans that are integrated (Fig. 3).
In summary, we have demonstrated that the contribution of gluconeogenesis to glucose production can be measured using only ~0.05% 2H in body water. This requires that one take advantage of the fact that isotope fractionation occurs during gas chromatography; we demonstrated that biasing the ion chromatogram integration method and quantifying the ion current ratio across the leading edge of the chromatographic peaks enhance the detection of 2H. We have discussed how the ability to use a low dose of 2H2O may be advantageous in certain situations. We hope that our approach to integrating ion chromatograms will facilitate studies of metabolism in which other 2H-labeled tracers are used. Future directions of our research are aimed at modulating parameters that enhance isotope fractionation and applying this method in studies using other 2H-labeled tracers.
![]() |
ACKNOWLEDGEMENTS |
---|
This project was supported by the Mt. Sinai Health Care Foundation of Cleveland and the Diabetes Association of Greater Cleveland (Grant no. 449-01). V. Chandramouli was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-14507. V. E. Anderson was supported by National Science Foundation Grant MCB-0110610.
![]() |
FOOTNOTES |
---|
Address for reprint requests and other correspondence: S. F. Previs, Dept. of Nutrition, Dental Bldg., Rm. 201, Case Western Reserve Univ. School of Medicine, 10900 Euclid Ave., Cleveland, OH 44106-4906 (E-mail: sxp29{at}po.cwru.edu).
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.
First published February 11, 2003;10.1152/ajpendo.00485.2002
Received 7 November 2002; accepted in final form 3 February 2003.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Babcock, MB,
and
Cardell RR.
Hepatic glycogen patterns in fasted and fed rats.
Am J Anat
140:
299-338,
1974[ISI][Medline].
2.
Boden, G,
Chen X,
and
Stein TP.
Gluconeogenesis in moderately and severely hyperglycemic patients with type 2 diabetes mellitus.
Am J Physiol Endocrinol Metab
280:
E23-E30,
2001
3.
Chandramouli, V,
Ekberg K,
Schumann WC,
Kalhan SC,
Wahren J,
and
Landau BR.
Quantifying gluconeogenesis during fasting.
Am J Physiol Endocrinol Metab
273:
E1209-E1215,
1997
4.
Dekker, E,
Romijn JA,
Ekberg K,
Wahren J,
Van Thien H,
Ackermans MT,
Thuy LT,
Chandramouli V,
Kager PA,
Landau BR,
and
Sauerwein HP.
Glucose production and gluconeogenesis in adults with uncomplicated falciparum malaria.
Am J Physiol Endocrinol Metab
272:
E1059-E1064,
1997
5.
Gastaldelli, A,
Baldi S,
Pettiti M,
Toschi E,
Camastra S,
Natali A,
Landau BR,
and
Ferrannini E.
Influence of obesity and type 2 diabetes on gluconeogenesis and glucose output in humans: a quantitative study.
Diabetes
49:
1367-1373,
2000[Abstract].
6.
Gastaldelli, A,
Toschi E,
Pettiti M,
Frascerra S,
Quinones-Galvan A,
Sironi AM,
Natali A,
and
Ferrannini E.
Effect of physiological hyperinsulinemia on gluconeogenesis in nondiabetic subjects and in type 2 diabetic patients.
Diabetes
50:
1807-1812,
2001
7.
Jones, JG,
Solomon MA,
Cole SM,
Sherry AD,
and
Malloy CR.
An integrated 2H and 13C NMR study of gluconeogenesis and TCA cycle flux in humans.
Am J Physiol Endocrinol Metab
28:
E848-E856,
2001.
8.
Kalhan, SC,
Parimi P,
Van Beek R,
Gilfillan C,
Saker F,
Gruca L,
and
Sauer PJ.
Estimation of gluconeogenesis in newborn infants.
Am J Physiol Endocrinol Metab
281:
E991-E997,
2001
9.
Landau, BR.
Quantifying the contribution of gluconeogenesis to glucose production in fasted human subjects using stable isotopes.
Proc Nutr Soc
58:
963-972,
1999[ISI][Medline].
10.
Matthews, DE,
and
Hayes JM.
Systematic errors in gas chromatography-mass spectrometry isotope ratio measurements.
Anal Chem
48:
1375-1382,
1975[ISI].
11.
Patterson, BW.
Determination of amino acid isotopic enrichment: methods, difficulties and calculations.
In: Methods for Investigation of Amino Acid and Protein Metabolism, edited by El-Khoury AE.. Boca Raton, FL: CRC, 1999, p. 103-119.
12.
Petersen, KF,
Krssak M,
Navarro V,
Chandramouli V,
Hundal R,
Schumann WC,
Landau BR,
and
Shulman GI.
Contributions of net hepatic glycogenolysis and gluconeogenesis to glucose production in cirrhosis.
Am J Physiol Endocrinol Metab
276:
E529-E535,
1999
13.
Postle, AD,
and
Bloxham DP.
The use of tritiated water to measure absolute rates of hepatic glycogen synthesis.
Biochem J
192:
65-73,
1980[ISI][Medline].
14.
Previs, SF,
and
Bunengraber H.
Methods for measuring gluconeogenesis in vivo.
Curr Opin Clin Nutr Metab Care
1:
461-465,
1998[Medline].
15.
Rothman, DL,
Magnusson I,
Katz LD,
Shulman RG,
and
Shulman GI.
Quantitation of hepatic glycogenolysis and gluconeogenesis in fasting humans with 13C NMR.
Science
254:
573-576,
1991[ISI][Medline].
16.
Schumann, WC,
Gastaldelli A,
Chandramouli V,
Previs SF,
Pettiti M,
Ferrannini E,
and
Landau BR.
Determination of the enrichment of the hydrogen bound to carbon 5 of glucose on 2H2O administration.
Anal Biochem
297:
195-197,
2001[ISI][Medline].
17.
Thomas, LC,
and
Weichman W.
Quantitative measurements via co-elution and dual-isotope detection by gas chromatography-mass spectrometry.
J Chromatogr
587:
255-262,
1991[Medline].
18.
Yang, D,
Diraison F,
Beylot M,
Brunengraber DZ,
Samols MA,
Anderson VE,
and
Brunengraber H.
Assay of low 2H enrichment by isotopic exchange with [U-13C3]acetone and gas chromatography-mass spectrometry.
Anal Biochem
258:
315-321,
1998[ISI][Medline].