ABSTRACT
The glycemic index (GI) measures the magnitude of the postprandial increase in
blood glucose caused by a test food compared with a reference food/beverage, such as a
glucose solution or white bread, containing the same amount of carbohydrate. The insulin index is determined in a similar manner of GI calculation, except that blood insulin AUC is used in place of blood glucose AUC. Low GI and insulin index foods are desirable because
foods with low GI and insulin index result in gradual increase in postprandial
glycemia/insulinemia and lower blood glucose/insulin fluctuations compared with foods
with high GI and insulin index. This attenuated glycemic and insulinemic responses of
low GI and insulin index foods is associated with reduced risks of obesity, diabetes
mellitus, and chronic diseases.
Fructose has low glycemic and insulin index, and a high fructose-containing food, such
as raisins, would be expected to have low glycemic and insulin index values. In addition
to its low glycemic and insulin index, small, or “catalytic” amounts of fructose (e.g., 5-
10 g) lower the glycemic response to other carbohydrates. However, prefeeding of fructose is necessary to achieve this effect due to the slow intestinal absorption of fructose. The overall objective of this dissertation was to investigate the current interest of carbohydrate metabolism. The first goal was to determine a difference in carbohydrate
iii metabolism in populations with different metabolic status. The second objective was to
determine fructose absorption in the presence of erythritol in vivo and in vitro.
The GI and insulin index of raisins were determined and compared in healthy sedentary
young adults, endurance athletes, and people with impaired glucose tolerance. The GI of
raisins was low (GI ≤55) in the healthy sedentary people (49.4 ± 7.4) and people with
pre-diabetes (49.6 ± 4.8) and was moderate (GI 55-69) in the athletes (62.3 ± 10.5), but
there were no differences among the subject groups (P = 0.437). The insulin index of raisins was not significantly different among the groups. Raisins are a low to moderate GI food, with a correspondingly low insulin index.
In the second study, the effect of the simultaneous ingestion of an equimolar
amount of erythritol and fructose on fructose absorption was determined in healthy
subjects. Breath hydrogen production with a beverage of equimolar mixture of 50 g
fructose and 33.3 g erythritol (FE) was 207% higher than that of a beverage of 50 g
fructose (F) (P <0.05). Serum fructose levels were 20% lower in the FE compared with F
(P < 0.05). However, serum erythritol level in FE was increased to 727.7 ± 22.2
mmol/min/L, which it was negligible in F (P < 0.05). The rise in breath hydrogen levels
in FE versus F indicated greater carbohydrate malabsorption. Because of the considerable
rise in serum erythritol and the decrease in serum fructose in the FE versus F groups, it
appeared that erythritol was absorbed at the expense of fructose.
The third study investigated the effect of erythritol on intestinal fructose
absorption using Caco-2 cells. The inhibitory effect of erythritol on fructose absorption
that we observed in healthy humans was reproducible in a Caco-2 cell model at high
iii doses of fructose and erythritol. Erythritol inhibited fructose absorption in a dose-
dependent fashion (P < 0.05).
In conclusion, raisins, a high fructose-containing food, has a low GI and insulin
index. A mixture of an equimolar amount of fructose and erythritol also had low
glycemic and insulinemic responses. However, increased gastrointestinal distress after intake of the mixture of fructose and erythritol may limit the usage of those carbohydrates.
Further research should focus on the interaction of sugar alcohol s and other food components which may impact on carbohydrate absorption and gastrointestinal distress.
iv
Dedicated to my Lord, Jesus Christ
v ACKNOWLEDGMENTS
It has been a quite journey with joy and tears, and this dissertation would not have been possible without support, encouragement, and prayer of these people. I specially appreciate Dr. Steve Hertzler. His invaluable support, intellectual idea, guidance, and expertise have facilitated my growth as a scientist. Especially, I thank him for his patience and deep understanding of the most difficult period of my study. I am grateful to Dr. Anne Smith for her advice, guidance, and kindness. Her invaluable support and encouragement was crucial for the completion of my study. I thank Dr. Mark Failla for his help, discussion, advice, and expertise with cell culture model. I also wish to thank my committee member, Dr. Martha Belury and Dr. Gail Kaye, for great suggestions and advice. I appreciate Dr. Sonhee Park for her advice, encouragement, friendship, and prayer all through my difficult days. I thank Bryan Wolf, Jennifer Williams, and Courtney Colombo for the opportunity to work with them and learn from them. I would like to thank Michelle Asp, Rubina Khan, and other study assistants for their hard work. I thank all the subjects who participated in my two clinical trials for their endurance, cooperation, and kindness. I wish to thank to Tianyao Huo and Sagar Thakkar for their suggestions and technical support regarding cell culture. I acknowledge the support and love from my family during my study. I thank concerns, encouragements, and prayer of my friends. The research was supported by Abbott Laboratories, Ross Products Division and California Raisins Marketing Board.
vi VITA
November 15, 1973……………………….…………………….Born-Seoul, South Korea 1996……….………………………..………...………………….B.S. Food and Nutrition Ewha Womans University 1998……....…………………………………………...………….M.S. Food and Nutrition Ewha Womans University 1999-2006……………………………………...... Graduate Research Assistant The Ohio State University 2005……………………………………………………….....Graduate Teaching Assistant The Ohio State University 2006-Present………………………………………………………………...Dietetic Intern The Ohio State University
PUBLICATIONS
1. Hertzler S, Kim Y, Khan R, Asp M, Savaiano D. (2006) Intestinal disaccharidase depletions, In: Shils ME, Shike M, Ross AC, Cabellero B, Cousins RJ, eds. Modern Nutrition in Health and Disease. 10th ed. Philadelphia, PA: Lippincott Williams & Wilkins;189-1200.
2. Hertzler SR and Kim Y. Glycemic and insulinemic responses to energy bars of differing macronutrient composition in healthy adults. Med Sci Monit 2003; 9(2):CR84- 90.
3. Chang, N., Kim, K., and Kim, Y. Folate nutritional status of women of childbearing age. Nutritonal Sciences 1999; 2(1):51-55
vii 4. Kim, Y., Kim, K., and Chang, N. Dietary folate intake of Korean women of childbearing age. Korean J. Nutriton 1999; 32(5):585-591
5. Chang, N., Kim, Y., and Kim, K. Effects of dietary folate concentrations on plasma and tissue folate concentrations in rats. Korean J. Nutrition 1998; 31(3):243-252.
6. Chang, N., Kim, K., Kim, Y., Seo JB, and Kwon O. Effects of alcohol administration and dietary folate on plasma homocysteine and liver histopathology. Korean J. Nutrition 1998; 31(7):1121-1129
FIELDS OF STUDY
Major Field: Nutrition
viii TABLE OF CONTENTS
Page
Abstract…………………………………………………………………………………..ii
Dedication……………………………………………………………………………….v
Acknowledgments……………………………………………………………………….vi
Vita……………………………………………………………………………………...vii
List of Tables………………………………………………………………………...…xii
List of Figures………………………………………………………………………….xiii
Chapters:
1. Introduction………………………………………………………………………....1
2. Literature Review…………………………………………………………………...8 Glycemic index……………………………………………………………………..8 Methodological aspects of glycemic index testing protocols…………………..9 Food factors influencing glycemic index……………………………………..12 Subject characteristics that influence glycemic index………………………...15 Insulin index………………………………………………………………….18 Catalytic doses of fructose, glucokinase, and postprandial glycemia……………..19 Fructose absorption……………………………………………………………….23 Clinical assessment of fructose absorption: breath hydrogen testing…………25 Fructose absorption capacity in humans………………………………………27 Dietary factors affecting fructose absorption………………………………...28 Glucose…………………………………………………………………...28 Amino acids………………………………………………………………31 Fructose, sorbitol, and carbohydrate malabsorption……………………...31 Facilitation of fructose absorption by glucose: proposed mechanisms……………32 Disaccharidase-related transport system……………………………………...32 Solvent drag mediated paracellular pathway…………………………………34 Apical GLUT2………………………………………………………………..36 ix Erythritol-Introduction…………………………………..………………………..40 Erythritol absorption…………………………………...………………………….42 Erythritol distribution……………………………………………………………...44 Metabolism of malabsorbed erythritol by the colonic microflora…………………45 Effect of erythritol on glycemic and insulinemia………………………………….47 Laxative effect of erythritol………………………………………………………..47 References………………...……………………………………………………….48
3. Determination of the glycemic and insulin index values of raisins in three populations………………………………………………………………….……..61 Introduction……………………………………………………………………..…61 Methods……………………………………………………………………..……..63 Results………………………………………………………………………..…....66 Discussion…………………………………………………………………..……..68 References…………………………………………………………………….…...73
4. Inhibition of fructose absorption by erythritol in healthy adults……..…..……….81 Introduction………………………………………………………………………..81 Methods…………………………………………………………………………....83 Results……………………………………………………………………………..87 Discussion…………………………………………………………………………90 References…………………………………………………………………………94
5. Erythritol inhibits transepithelial transport of fructose across Caco-2 human intestinal cells…………………………………………………………………….108 Introduction…………………………………………………………...………….108 Methods…………………………………………………………………………..110 Results…………..………………………………………………………………..115 Discussion………………………………………………………………………..117 References………………………………………………………………………..121
6. Conclusions………………………………………………………………………129
Bibliography…………………………………………………………………….…….132
Appendix
A. Institutional review board approval letter for the raisin study………………..152
B. Recruitment flyer for the raisin study…………………………………………154
C. Screening questionnaire for the raisin study…………………………………..156
D. Treatment visit data collection form for the raisin study……………………..163
x E. Western institutional review board approval letter for the erythritol and fructose study………………………………………………………………….168
F. Recruitment flyer for the erythritol and fructose study…………………….….171
G. Screening questionnaire for the erythritol and fructose study………………...173
H. Breath hydrogen and fasting plasma glucose screening form for the erythritol and fructose study……………………………………………………………..179
I. Test visit data collection for the erythritol and fructose study…………………181
J. Flatus/bowel movement form for the erythritol and fructose study…………...187
K. Gastrointestinal tolerance factors form for the erythritol and fructose study…189
L. Exit visit questionnaire for the erythritol and fructose study…………………………………………………………………………...191
xi LIST OF TABLES
Table Page
2.1. Fructose absorption capability studies in healthy humans………………………..30
2.2. Carbohydrate composition of fruit juices (g/100 mL serving)…………………....32
2.3. Concentrations of erythritol in various foods……………………………………..41
3.1. Positive incremental areas under the curve (IAUC), excursion, and baseline- adjusted peak (BAP) for serum glucose and corresponding glycemic index of raisins in the S group (N=10), the A group (N=11) and the P group (N=10)…...... 79
3.2. Positive incremental areas under the curve (IAUC), excursion, and baseline- adjusted peak (BAP) for serum insulin and corresponding glycemic index of raisins in the S group (N=10), the A group (N=11) and the P group (N=10). ……….…..80
4.1. Postprandial breath hydrogen, serum fructose, erythritol, glucose, insulin, and lactate areas under the curve (AUC) in response all study beverages…………...106
4.2. Self-reported frequencies of rectal gas passages and bowel movements and stool consistency ratings in the postprandial period in response to each study beverage. ……………………………………………………………………………………107
xii LIST OF FIGURES
Figure Page
2.1. Glucokinase regulation in the liver……………………………………………………22
2.2 Fructose absorption in the small intestine……………………………………………..24
2.3. Schematic illustration of the trans-and para-cellular pathway in the epithelial cells. ……………………………………………………………………………………..35
2.4. The apical GLUT2 model of glucose absorption (A) before a meal and (B) after a meal ……………………………………………………………………………..38
2.5. Chemical structure of erythritol……………………………….…………………..40
3.1. Serum glucose and insulin responses to raisins and glucose solution in sedentary (Group S), athletically trained (Group S), and pre-diabetic (Group P) subjects…..79
4.1. Postprandial breath hydrogen concentrations for all beverages at each time oint.100
4.2. Postprandial serum fructose concentrations for all beverages at each time point..101
4.3. Postprandial serum erythritol concentrations at each time point………………...102
4.4. Postprandial serum glucose centrations for all beverages at each time point…....103
4.5. Postprandial serum insulin concentrations for all beverages at each time point…104
4.6. Postprandial serum lactate concentrations for all beverages at each time point…105
5.1. Experimental design……………………………………………………………...111
5.2. Transport of fructose into the basolateral compartment for different length of incubation……………………………………………………………………...…124
5.3. Transport of fructose and erythritol into the basolateral compartment at high doses of fructose and erythritol…………………………………………………………125
xiii 5.4. Transport of fructose and erythritol into the basolateral compartment after phenol red assay was conducted the day before the xperiment…………………….…..126
5.5 Transport of fructose and erythritol into the basolateral compartment after apical side was pre-treated with glucose-free DMEM the day before the xperiment….127
5.6 Transport of fructose and erythritol to the basolateral compartment at low doses of fructose and erythritol……………………………………………………..…128
xiv CHAPTER 1
INTRODUCTION
1.1. Background
The glycemic index (GI) is a physiologic carbohydrate classification system
based on the magnitude of the increase in blood glucose caused by a particular food
compared with the rise observed for a reference food such as a glucose solution or white
bread (1). The GI concept suggests that carbohydrates with a low GI result in a more
moderate, sustained increase in postprandial glycemia and a lower peak blood glucose
level than carbohydrates with a high GI (2). Low glycemic index foods have been
recommended in weight management, dietary management of diabetes, and sports nutrition (3). In addition, the glycemic index is used as a means to evaluate carbohydrate
digestibility because slowly digested carbohydrates produce low GI values (4). The
insulin index is a numerical index that ranks carbohydrate based on their rate of insulin
response. High fasting and/or postprandial insulin levels due to a high insulin index diet
are associated with an increased risk of insulin resistance and chronic diseases such as
obesity, cardiovascular disease (Frost et al 1999, Salmeron et al 1997). Although GIs of
most of foods are positively correlated with the insulin index of the foods, protein-rich
foods cause higher insulin responses that expected from their GI because protein induces
1 insulin secretion. Thus, it is increasingly recognized that the knowledge of the GI of a
food, while important, is not complete without the knowledge of the insulin response as
well.
Fructose has a low GI (the GI of fructose is 19 where glucose = 100) (5) and insulin index (the insulin index of fructose is 22 where white bread = 100) (6). Because
about 50% of the carbohydrate content of raisins is fructose (7), it might be expected that
raisins would be a low GI food. There is only one previous study (1), in which the GI of
raisins was measured in healthy subjects. In this study, the GI of raisins was 64, where the GI
of glucose = 100 (a moderate GI). However, this study was published ~25 years ago, had a
small sample size (n = 6), and no data on postprandial insulin secretion were obtained.
Further, it is not known if the GI of raisins remains consistent when measured in subjects with different levels of habitual physical activity or alterations in glucose metabolism.
Thus, the GI and insulin index values of raisins were determined in a larger group of
subjects. In addition, the effects of habitually high levels of physical activity and also impaired fasting glucose on the measurement of the GI of raisins were examined.
Not only does fructose have a low GI, but small doses (referred to as “catalytic”
doses) may also lower the glycemic response to other carbohydrates in the meal (8). It
was demonstrated in a previous study from our laboratory that 10 g fructose consumed
30-60 min before feeding decreased the blood glucose response to a starchy carbohydrate
(mashed potatoes) (8), an effect that may be of benefit to people with diabetes. However,
in this study, administration of fructose simultaneously with the mashed potatoes did not
reduce the glycemic response. Slow intestinal absorption of fructose is a possible
2 explanation for these results. At least 60% of healthy adults will malabsorb a 50 g
fructose dose, with some subjects absorbing as little as 5-10 g (9, 10).
Free glucose has a well-known facilitating effect on fructose absorption from the
intestine (11, 12), but the addition of free glucose to a carbohydrate-containing food
would add calories and increase glycemia. The discovery of a non-caloric, non-glycemic
system for enhancing the intestinal absorption of fructose would be desirable to take
advantage of fructose’s ability to lower postprandial glycemia.
Erythritol, a 4-carbon sugar alcohol, is hypothesized to be a good candidate for facilitating the absorption of fructose. Erythritol is very well absorbed from the intestine, presumably via a paracellular pathway, and there is evidence that erythritol enhances the intestinal absorption of calcium by this route (13). Because the facilitation of fructose absorption by glucose is thought to occur paracellularly (11, 14), it was hypothesized that erythritol might have a similar effect. Erythritol also has an advantage over glucose with regard to blood glucose control because it is both noncaloric and nonglycemic (15). Therefore, two studies were conducted to examine the effects of
erthritol on the absorption of fructose in humans and Caco-2 cells.
1.1. Objectives/specific aims
The overall objective of this dissertation was to investigate the current interest of
carbohydrate metabolism. The first goal was to determine a difference in carbohydrate
metabolism in populations with different metabolic status. The second objective was to
determine fructose absorption in the presence of erythritol.
For the study described in Chapter 3, the goal was to measure the glycemic and
3 insulin index of raisins in 3 different populations: healthy sedentary young adults,
competitive endurance athletes, and people with impaired fasting glucose. The objective of the study in Chapter 4 was to determine if the simultaneous ingestion of an equimolar amount of erythritol affects fructose absorption in healthy adults. The purpose of the project in Chapter 5 was to investigate the effects of erythritol on fructose absorption at
the cellular level in a Caco-2 cell model that simulates human small intestinal cells.
1.3. Hypotheses
The following null hypotheses will be tested:
Chapter 3
• The GI of raisins will be similar to the previously published GI value of 64
• The GI value of raisins will be consistent, regardless of the subject population in
which the GI is determined
• The insulin index value of raisins will be consistent, regardless of the subject
population in which the insulin index is determined
Chapter 4
• Fructose absorption in healthy subjects will not be affected by the simultaneous
ingestion of an equimolar amount of erythritol in solution when compared with
the ingestion of fructose alone.
• Breath hydrogen AUC (an indicator of carbohydrate malabsorption) of a beverage
of equimolar mixture of fructose and erythritol will be same with a beverage of
fructose only.
• Serum fructose AUC will be unchanged in a beverage of equimolar mixture of 4 fructose and erythritol versus a beverage of fructose.
Chapter 5
• Fructose transport in Caco-2 cells will be unchanged by the presence of equimolar
amount of erythritol compared with the presence of fructose alone.
1.4. Significance of studies
The significance of the study in Chapter 3 is that it provides an updated GI value
for raisins, using a standardized sample size, and contributes data regarding the insulin
response as well. In addition, this study will address the question of whether the GI value of a food is consistent without regard to the physical activity level or glucose tolerance of
the subjects upon which the GI is determined.
The studies in Chapters 4 and 5 contribute valuable information regarding potential interactions between erythritol and fructose during intestinal absorption. This is important for food manufacturers who may wish to use either fructose or erythritol as a way of decreasing the glycemic response to their products while, at the same time, minimizing the risk of gastrointestinal distress due to carbohydrate malabsorption.
References
1. Jenkins DJA, Wolever TMS, Taylor RH, Barker H, Fielden H, Baldwin JM, et al.
Glycemic index of foods: a physiological basis for carbohydrate exchange. Am J
Clin Nutr. 1981;34:362-366.
2. Jenkins DJA, Kendall CWC, Augustin LSA, Franceschi S, Hamidi M, Marchie A,
5 et al. Glycemic index: overview of implications in health and disease. Am J Clin
Nutr. 2002;76(suppl):266S-273S.
3. FAO food and nutrition paper 66. Carbohydrates in human nutrition. Report of an
FAO/WHO expert consultation on carbohydrates, April 14-18, 1997, Rome, Italy.
Rome:FAO;1998
4. Keim NL, Levin RJ, Havel PJ. Carbohydrates. In: Shils ME, Shike M, Ross AC,
Cabellero B, Cousins RJ, eds. Modern Nutrition in Health and Disease. 10th ed.
Philadelphia, PA: Lippincott Williams & Wilkins;p78
5. Foster-Powell K, Holt SHA, Brand-Miller JC. International table of glycemic
index and glycemc load values: 2002. Am J Clin Nutr. 2002;76:5-56.
6. Lee BM, Wolever TMS. Effect of glucose, sucrose, and fructose on plasma
glucose and insulin responses in normal humans: comparison with white bread.
Eur J Clin Nutr. 1998; 52: 924-928.
7. Matthews RH, Pehrsson PR, and Farhat-Sabet M. Sugar content of selected foods:
individual and total sugars. United States Department of Agriculture. Human
Nutrition Information Service. Home Economics Research Report Number 48.
1987.
8. Heacock PM, Hertzler SR, Wolf BW. Fructose prefeeding reduces the glycemic
response to a high-glycemic index, starchy food in humans. J Nutr. 2002;
132:2601-2604.
9. Rumessen JJ and Gudmand-Hoyer E. Absorption capacity of fructose in healthy
adults. Comparison with sucrose and its constituent monosaccharides.
Gut.1986;27:1161-1168.
6 10. Trustwell AS, Seach JM, Thorburn AW. Incomplete absorption of pure fructose
in healthy subjects and the facilicating effect of glucose. Am J Clin Nutr.
1988;48:1424-1430.
11. Shi X, Schedl HP, Summers RM, et al. Fructose transport mechanisms in humans.
Gastroenterol. 1997;113:1171-1179.
12. Fujisawa T, Riby J, Kretchmer N. Intestinal absorption of fructose in the rat.
Gastroenterol. 1991;101: 360-367.
13. Mineo H, Hara H, Tomita F. Sugar alcohols enhance calcium transport from rat
small and large intestine epithelium In Vitro. Dig Dis Sci. 2002;47(6):1326-1333.
14. Turner JR, Cohen DE, Mrsny RJ, Madara JL. Noninvasive in vivo analysis of
human small intestinal paracellular absorption: regulation by Na+-glucose
cotransport. Dig Dis Sci. 2000;45(11):2122-2126.
15. Ishikawa M, Miyashita M, Kawashima Y, et al. Effects of oral administration of
erythritol on patients with diabetes. Regulatory toxicology and pharmacology.
1996;24:S303-S308.
7 CHAPTER 2
LITERATURE REVIEW
Glycemic index
The glycemic index (GI) is a physiologic carbohydrate classification system based on the magnitude of the postprandial increase in blood glucose caused by a particular food (1). Wolever (2, 3) defines the GI of a test food as:
Positive incremental area under the blood glucose curve for the test food (50 g available carbohydrate) GI = X 100 Positive incremental area under the blood glucose curve for the reference food (white bread or glucose solution, 50 g available carbohydrate)
A glucose solution (50 g) was initially used as the reference, but subsequent studies employed white bread because it is a common food in the diet and more physiologic than the glucose solution (2-4). However, the nutritional composition of white bread is variable, possibly producing diverse GI values for the same foods when tested by different laboratories (2). Because it is easier to standardize from one laboratory to the
8 next, the glucose solution is generally preferred as a reference. However, there may be situations in which a different reference (e.g., white bread or rice) is desired. The GI
values of test foods obtained when white bread is used as a reference are 1.4 times those obtained when glucose solution is the reference (4). In Japan, white rice has been used as
a reference food due to availability and palatability of rice in Asian cultures (5).
A GI of 55 or less is low GI, a GI between 56 and 69 is medium, and a GI 70 or
more is high. The cutoff 70 for a high GI is because most of bread, breakfast cereals, and
potatoes have GI more than 70. These foods have low GI counterparts which are less than
55 (personal correspondence with Dr. J. Brand-Miller). The GI concept suggests that
carbohydrates with a low GI result in a more moderate and sustained increase in
postprandial glycemia and a lower peak blood glucose level than carbohydrates with a
high GI (6). There are many physiological and nonphysiological factors that influence
GI values. General classification of these factors include: 1) methodological aspects of
glycemic index testing protocols; 2) factors associated with the composition of the test
foods; and 3) physical characteristics of the test subjects. Some of these factors will be
reviewed in the next section.
Methodological aspects of GI testing protocols
One factor that has created much confusion in the interpretation of GI studies is
the method for determining the area under the postprandial blood glucose curve. The
accepted method for calculating area under the curve is the positive incremental area
under the curve (AUC), including only the area above the fasting value (3). However,
some laboratory groups have calculated total versus positive incremental AUC. The total
AUC includes all the area down to a blood glucose value of zero (rather than just down to
9 the baseline value). This approach dramatically reduces the differences in postprandial
glucose AUC between foods and the use of this unaccepted procedure has been
responsible for the erroneous conclusion that the GI concept is not applicable to mixed
meals (4, 7-9).
The method of blood sampling is another factor affecting the postprandial
glycemic response. Blood can be obtained from a vein, an artery, or a capillary bed.
Although collecting arterial blood is ideal for assessing the glycemic responses to foods, it is risky and uncomfortable to study subjects (3, 10). Capillary blood is preferred
because it is easy to obtain, its glucose level is very close to that found in arterial blood,
and the postprandial glucose response from capillary blood is less variable than that from
venous blood (2, 4, 9). An interlaboratory study was conducted in which GI of the same
5 foods (potatoes, rice, sphaghetti, and barley, and white bread, with glucose as a
reference) was measured in 7 different laboratories. Those laboratories in which venous
samples were collected had a larger coefficient of variation (CV) in the glucose reference
AUC (>50%), compared with 23 % CV in the laboratories in which capillary blood was
sampled (11). Although there was greater variability of GI values with venous versus
capillary blood in that study, the mean GI value was not affected. Because of the lower
variability with capillary samples versus venous samples, the sensitivity of detecting
significant differences among the GI values of different foods is enhanced (3).
Blood samples must be collected until the blood glucose level returns to at or near
the baseline value because the GI of a test food is calculated the area under the glycemic
index curve and above the baseline (2, 12). Therefore, the glucose response is typically
10 measured over 2 h in normal subjects (baseline, 15, 30, 45, 60, 90, and 120 minutes) and over 3 h in diabetic patients (12).
To determine how much available carbohydrate portion would be used for GI test, a portion of food containing 50 g of available carbohydrate (total carbohydrate minus dietary fiber) was suggested (1). However, for a food having low amount of carbohydrate per serving, an unrealistically large serving of the food would be consumed to obtain a 50-g carbohydrate load. For example, a medium-sized raw carrot has 5.1 g of available carbohydrate/72 g (13). To achieve a 50-g carbohydrate load, 705.9 g of carrots would have to be consumed. This is a very large amount of carrots and it may not be fully consumed by subjects in a limited time period. Thus, for foods such as carrots, a 25-g carbohydrate portion is used (along with a corresponding 25-g glucose solution as the reference). In a study in which different carbohydrate loads (25 g, 50 g, and 100 g) were evaluated, the “absolute” glycemic responses to different levels of carbohydrate in the same test food differed. Nevertheless, the “relative” glycemic responses (compared with a reference containing the same amount of carbohydrate) are the same at any level of carbohydrate (14, 15).
In preparation for GI tests, subjects are instructed not to smoke, drink alcoholic beverages, or exercise the day before the tests. Cigarette smoking, alcohol intake, and vigorous exercise may have confounding effects on GI tests (3). Cigarette smoking causes acute insulin resistance, so smoking should not be permitted on the day of the test.
Alcohol consumption inhibits gluconeogenesis (16), and may alter other factors such as the oxidation of macronutrients or insulin sensitivity (17). Acute physical exercise the day before the test increases muscle glucose uptake and insulin sensitivity (3). The meal
11 test is recommended to be fed at breakfast after 10-14 hour overnight fast because having
subjects in the fasted state minimizes between-subject variation (3). Wolever and
Bolognesi (18) measured the relative glycemic responses to two cereals in the morning
after 10-14 h overnight fast and at lunchtime after standard breakfast. The difference in
glycemic response between two cereals tested in the morning was significantly greater
than that conducted at lunchtime (103 ± 13 vs. 52 ± 6 mmol·min/L, respectively).
Food factors influencing GI
The rate of carbohydrate absorption in the small intestine plays an important role
in determining the metabolic effects of dietary carbohydrate. Food factors that affect the
carbohydrate absorption rate include the physical and chemical structures of starches and
sugars, the presence of protein and fat, cooking methods used, the ripeness of fruits, and
the presence of anti-nutrients (19).
Different monosaccharides have different GI values. The GI of monosaccharides
was measured in young healthy subjects, with white bread used as the reference food (1).
The GI of glucose was 136.6 ± 22.0, and that of fructose was 16.4 ± 5.5. These authors
also tested sucrose, which had an intermediate GI (83.3 ± 14.9). This is likely due to the presence of equimolar amounts of glucose and fructose that appear in the intestinal lumen
upon sucrose digestion.
The GI of fruit juices is largely dependent on the absorption of the constituent
carbohydrates. The GI of apple juice is 40 (where glycemic index of glucose is 100) (20),
which is considered to be a low GI (GI < 55 is low). Apple juice has more than twice as
much fructose as glucose and contains sorbitol (a poorly absorbed sugar alcohol). In
contrast to apple juice, orange juice has equimolar amount of fructose (2.4 g / 100 mL
12 orange juice) and glucose (2.4 g / 100 mL orange juice) and no sorbitol, which resulted in
no gastrointestinal symptoms and more complete carbohydrate absorption (21), yielding
a GI of 52 (20), higher than the GI of apple juice. The lower GI value of apple juice
compared with orange juice might be due to the higher ratio of fructose to glucose and
the presence of sorbitol.
Starch characteristics also affect GI values. For example, white low-amylose rice
varieties have a higher GI (GI = 83 – 139, where GI glucose = 100) compared with that
of high-amylose types of rice (GI = 37-64) (20). Amylose is not readily broken down by
amylase as compared with amylopectin. Hence, amylose is slowly digested, producing
low GI (5).
Addition of protein and fat to carbohydrate foods affects glycemic response by delaying gastric emptying and stimulating insulin secretion (22, 23). Spiller et al.
determined the effect of various amounts of protein on serum glucose and insulin
responses in healthy subjects (24). Subjects were administered test beverages containing
0, 15.8, 25.1, 33.6, or 49.9 g protein together with 58 g carbohydrate. Protein-containing
beverages caused significantly lower blood glucose AUC than the protein-free beverage
(P < 0.01), and the effect of protein on lowering blood glucose response was linear
(r=0.986, P < 0.001). Administration of the various amount of protein resulted in an
increase in insulin AUC (P < 0.01), but it was not dose-dependent.
It is generally believed that fat reduces postprandial blood glucose because fat
reduces gastric-emptying rate caused by increased secretion of glucose-dependent-
insulin-releasing polypeptide (GIP) and glucagons-like polypeptide-1 (GLP-1) (25). In a
study to determine the effect of fat on glucose response, healthy subjects consumed 50 g
13 available carbohydrate (white bread) along with 0, 5, 10, 20, or 40 g fat (non-
hydrogenated-fat margarine). Intake of 40 g fat produced 30 % reduced incremental AUC
of blood glucose compared with fat-free meal (P < 0.05). However, no different
incremental AUC was observed between meals containing 5, 10, and 20 g fat (25). On the
country, adding fat to potato has no effect on postprandial glucose responses in people
with type 2 diabetes (26), possibly due to an inability of fat to stimulate GLP-1 secretion.
Recently, Moghaddam et al. measured the effects of protein and fat on the glycemic
responses in nondiabetic hyperinsulinemic subjects (fasting plasma insulin ≥ 40 pmol/L)
and normal subjects (27). Subjects were administered 50 g glucose with 0, 5, 10, or 30 g
fat (corn oil) and/or 0, 5, 10, or 30 g protein (soy protein concentrate) dissolved in 250
mL water, and 3-d diet records were used to estimate dietary intake. Protein reduced the
glucose responses 2 times more than fat (P < 0.001), but there was no significant fat and protein interaction. Protein reduced glycemic responses to a greater extent in subjects with higher waist circumference (r = -0.56 for protein effect on glycemia versus waist circumference, P = 0.011) and a high intake of dietary fiber (r = -0.60 for dietery fiber effect on glycemia versus waist circumference, P = 0.005). However, these relationships were not observed in hyperinsulinemic subjects. In contrast, fat had the greater effects on glycemia in hyperinsulinemic subjects (r = 0.49 for fat effect on glycemia versus fasting insulin, P = 0.029) and was not affected by waist circumference or dietary fiber.
Therefore, it seems that the effect of fat and protein on glycemic responses is dependent on subjects’ characteristics.
Cooking methods also affect GI values. Boiled red potatoes consumed hot had
high GI of 89, whereas boiled red potatoes consumed cold had intermediate GI of 56 (P <
14 0.05) (28). Similarly, baked US Russet potatoes had significantly higher blood glucose
AUC when eaten after cooking compared with when they were refrigerated and reheated
(145 ± 16 vs. 101 ± 11 mmol·min/L, P<0.05) (28). During cooking, starch structure is
broken down, resulting in rapid hydrolysis by amylase. However, after cooling, starch
retrogrades and forms an irregular structure that is resistant to digestion. Repeated
cooling and heating produces more resistant starch, which would delay digestion and
absorption, thus producing low glycemic response (28). In previous studies, the GI of
potatoes reported various. Boiled Australian potatoes had high GI, ranging from 87 to
101 (29). In contrast, GI of boiled Canadian potatoes was reported to be 61 (28), which is
lower than Australian potatoes. The difference may be due to different varieties of
potatoes used and different cooking method. Potatoes in Fernandes et al. study (28) was
precooked, frozen, and reheated before intake, while they were consumed immediately
after cooking in Soh and Brand-Miller’s study (29).
The GI of a mixed meal can be predicted from the GI of its component
carbohydrates (8, 15, 30). Some researchers have argued that the GI concept does not
apply to mixed meals (7, 31), but these studies have used inappropriate methods for
calculating the postprandial glucose AUC (as mentioned previously). A mixed meal GI
can be calculated by first multiplying the GI of each carbohydrate source by the
proportion of carbohydrate coming from that source and then summing these results (3).
Subject characteristics that influence GI
In a GI study, the glycemic response to the test food is compared with the
glycemic response to a reference food (glucose solution or white bread) within the same
subject, an approach that should factor out much of the intra-individual variability in GI
15 measurements (12). As such, GI is not affected by subject characteristics such as age (11,
32), ethnicity (11), gender (11), and glucose tolerance status (8, 33, 34).
The effect of the age of the subjects on the GI for lentils was measured in 10 adults
(mean 46 y) and 7 children (mean 11 y) with type 1 diabetes (32). The GI of lentils was very similar between the children (43 ± 7) and the adults (44 ± 10). In the interlaboratory study mentioned previously, although age (mean range from 21.0 to 36.3 y) and ethnicity of subjects were significantly different among the sites, the GI of foods was not significantly related with age or ethnicity. In addition, GI was not associated with the gender and BMI of the subjects (11).
The concept of GI of foods applies to subjects regardless of their glucose tolerance status. The GI of cornflakes measured in 6 healthy subjects (109 ± 11, where GI of white bread = 100) was not significantly different from that in 5 patients with type 2 diabetes (123 ± 7) (33). The GI of potatoes also was not significantly different between the subjects, although it appears that healthy subjects had a somewhat higher GI (100 ±
12) for potatoes compared with diabetic subjects (73 ± 11). However, a key limitation of this study is the small number of subjects. It is preferred to measure GI of foods at least in 10 subjects (3). In one study, the GI values of 15 foods in diabetic patients were linearly related to those GI values obtained in healthy subjects (r=0.753, P<0.01) (33).
Several studies reported that GI of foods was the same regardless of insulin sensitivity.
The GI of spaghetti in Type 2 diabetes was similar with that in type 1 diabetic patients
(49 ± 7 vs 57 ± 8) (33). Similarly, the GI of rice was not significantly different between patients with type 1 and type 2 diabetes (14). In a study measuring the GI of 37 foods in
16 type 1 and type 2 diabetic subjects, no differences between subject groups were reported
(34).
GI of a test food is not supposed to be affected by subject characteristics, such as
training status or glucose tolerance status because the glycemic response to a test food is
compared with the glycemic response to a standard food within the same subject.
However, recently, two studies reported the possibility of the influence of the training
status of subjects on the GI measurement of breakfast cereals (35, 36). In the first study,
the GI values of 3 commercially available breakfast cereals were measured in 11 healthy
sedentary (mean 23.5 ± 3.5 y) and 12 endurance-trained men (mean 23.3 ± 3.3 y) with
similar anthropometric data. The endurance-trained subjects exercised at least four times
per week, whereas sedentary subjects were not involved with any exercise activity.
Capillary blood glucose incremental AUC for the reference food (glucose solution) was
almost identical for sedentary and endurance- trained subjects (208 ± 15 vs 202 ± 17
mmol·min·L-1, respectively) (35). Among the 3 breakfast cereals in the Mettler et al. (35), one cereal (Kellogg’s Special K®) had a significantly lower GI (difference of 25 units) in
endurance-trained subjects compared with sedentary people (P = 0.002). Endurance-
trained subjects had slightly, but not significantly, higher GI values for the other cereals
(P = 0.204 for Bio-Birchermüesli®, and P = 0.052 for fit-Crisp®) than sedentary subjects.
In the other study (36), a group of moderately-trained men was included in addition to groups of healthy sedentary men and endurance-trained men to determine if
the effect of training status on the GI of a breakfast cereal (Kellogg’s Special K®) is dependent on the amount of exercise. Moderately-trained subjects exercised average 2.5 sessions per week, compared with 7.6 sessions per week for the endurance- trained group.
17 The GI of the breakfast cereal (Kellog’s Special K®), which had been shown to be
significantly different between sedentary and endurance-trained subjects in the previous
study, was measured. The GI obtained from capillary blood was significantly higher for
sedentary subjects (by 23 units) than for endurance trained subjects (P=0.02), consistent
with the findings of the previous study (35). The GI value of the cereal (measured in
capillary blood) for the moderately-trained men was between the other groups, and was
not different from either of the groups. However, the GI as determined from venous
blood sample was not significantly different among any of the groups, a finding that is
consistent with several other studies showing the relatively insensitivity of venous versus
capillary blood samples to detect differences in GI between foods (as reviewed in this
section). The insulin index was not different among the groups (P = 0.65). The results
from the studies conducted by Mettler et al. (35, 36) suggest that training status may be
the first subject-specific factor to influence GI values.
Insulin index
Most GI studies have not measured the postprandial insulin responses to test foods
and reference foods along with the glycemic responses to test foods and reference foods.
Insulin index is a measure to quantify the insulin response to a test food. Insulin index is
a manner similar to GI, except that insulin AUC is used in place of glucose AUC and, in
some studies, food portions are standardized by energy content rather than carbohydrate
content due to the effects of other macronutrients on the insulin response (37). Generally,
it is recognized that there is a linear relationship between GI and insulin index; high GI
foods also have a high insulin index. The proportionality of the glycemic and insulin
responses is true for most foods containing primarily carbohydrate (37, 38). However,
18 some foods yield higher insulin responses than expected from their GI. Protein-rich foods, for example, cause a disproportionately higher insulin response compared with their glycemic response because protein stimulates insulin secretion (37). In a study by
Ostman et al. (39), milk had insulin index of 90 ± 8 in 10 healthy subjects (aged 28-47 y) with normal BMI (23.4 ± 2.1 kg/m2), despite of low GI of 30 ± 4. In the same study,
lactic acid was reported as another factor to affect insulin index (39). At present, there is
much less known about the clinical significance of the postprandial insulin response and
the factors that affect it than for the GI. Certainly, the measurement of both blood
glucose and insulin in GI studies is desirable, but the measurement of insulin is not as
easy as glucose and adds considerable expense.
Glycemic load (GL) is a combination of the type (GI) and the amount of
carbohydrate consumed because blood glucose response is affected by not only the
quality of carbohydrate but also the quantity of carbohydrate (40). Glycemic load is
calculated as GI divided by 100 multiplied by its available carbohydrate (total
carbohydrates minus dietary fiber). Glycemic response can be controlled by consuming
low-GI foods and/or small intake of carbohydrates.
“Catalytic” doses of fructose, glucokinase, and postprandial glycemia
Recent discoveries have focused on potentially beneficial effects of small
amounts (5-10 g) of fructose for increasing hepatic glucose uptake and thereby decreasing postprandial glycemia (an effect referred to as the “catalytic” effect of fructose). Intraportal infusion of a small amount of fructose augmented hepatic glucose
uptake during a hyperglycemic hyperinsulinemic clamp in dogs (42). In another study by
19 Shiota et al. (43), 44.4 µmol glucose/kg/min was intraduodenally infused either with or without 2.22 µmol/kg/min fructose in dogs. The presence of fructose increased net hepatic glucose uptake (28 ± 5 µmol/kg/min) compared with glucose alone (7 ± 2
µmol/kg/min). The increased hepatic glucose uptake resulted in increased glycogen synthesis (68% vs 49%) with versus without fructose. In addition, arterial blood glucose levels during fructose infusion were half as much as when glucose was infused alone.
Arterial plasma insulin levels in the presence of fructose also followed the same trend as the blood glucose response.
Human studies also have demonstrated the beneficial effects of a catalytic amount of fructose on glucose and insulin response (44, 45). The effect of 7.5 g of fructose, co-
administered with a 75-g oral glucose challenge, on glucose tolerance was investigated in
healthy adults (44) and people with type 2 diabetes (45). A small amount of fructose
decreased the area under the curve (AUC) of plasma glucose by 19% in healthy people
(44). Although the AUC of plasma insulin with a small amount of fructose was not
significantly decreased, half of the subjects showed a reduced insulin response to the
intake of glucose when fed the catalytic dose of fructose (44). However, in subjects with
type 2 diabetes (45), the plasma insulin AUC was reduced by 21% in the presence of fructose and was accompanied by a fall of 14% in the plasma glucose AUC, implying
that enhanced glucose tolerance in the presence of small amounts of fructose was not
caused by improved insulin secretion (45). It appears that a catalytic amount of fructose
caused improved glucose tolerance by stimulating glucose uptake via glucokinase
activation (45).
20 It is interesting that, although a couple of studies have shown that concurrent
administration of 7.5 g of fructose with a 75 g glucose challenge significantly reduces the
glycemic response to the glucose load (44, 45), the same may not be true when fructose is
fed with a starchy food. Heacock et al. (46) demonstrated that the incremental AUC of blood glucose was reduced 25% and 27% only when 10 g of fructose was administered
60 or 30 min, respectively, before the intake of 50 g available carbohydrate from instant mashed potatoes. However, feeding the same dose of fructose immediately with the potatoes had no effect on postprandial glycemia. Further, a study by Wolf et al. (47) found no additional glucose-lowering benefit of a 5-g fructose dose when fed simultaneously with a starch/maltodextrin meal that included a source of guar gum.
Heacock et al. (46) postulated that although it is well-known that free glucose facilitates
fructose absorption from the intestinal lumen, the initial digestion of starch to mainly
oligosaccharides in the lumen would not have the same facilitating effect. Thus, some
degree of “lead time” was required to allow the fructose to be absorbed before the large
challenge dose of starchy carbohydrate was fed.
The mechanism by which catalytic doses of fructose lower glycemia is thought to
be increased activity of glucokinase, as described in Figure 2.1 (47, 48). Glucokinase is
involved in the phosphorylation of glucose to glucose-6-phosphate in the liver and is a
critical first step in hepatic glucose uptake from the portal blood. Glucokinase has a
relatively low affinity for glucose (Km 7-8 mM), and is not inhibited by concentrations of glucose 6-phosphate (48, 49). When glucose concentrations in the portal vein are very
high, glucose is phosphorylated by glucokinase. The liver then takes up glucose and then
either oxidizes it or stores it as glycogen, hence lowering the amount of glucose released
21 to the peripheral tissues. Small amounts of fructose stimulate glucokinase activity in
vitro (49). In the postabsorptive state, glucokinase is inactive because it is bound to a
regulatory protein within the nucleus of the hepatocyte. The binding of fructose-6-
phosphate to the regulatory protein keeps glucokinase from being released. However, the
presence of a small amount of fructose from the diet generates fructose-1-phosphate (via
the action of fructokinase). Fructose 1-phosphate can then bind to the glucokinase
regulatory protein, displacing fructose-6-phosphate in the process. Because of this
binding, glucokinase is released from the regulatory protein, and the enzyme translocates
to the cytosol, where it is active (47, 48). The increased activity of glucokinase can
increase hepatic uptake of glucose, thereby lowering postprandial glycemia.
Fructose Glucose Fructokinase
Fructose-1 -phosphate Glucose Glucose-6 - phosphate Inactive Active glucokinase glucokinase Fructose-6 - Fructose-6 - Fructose-6 - phosphate phosphate phosphate GKRP Nucleus GKRP glycolysis Cytosol
Figure 2.1. Glucokinase regulation in the liver. GKRP, glucokinsase regulatory protein. (47,
48)
Fructose absorption 22 Fructose absorption from the lumen of the small intestine into the blood is
achieved mainly via the enterocyte. The enterocyte is a polarized cell which consists of a
brush border membrane in contact with the lumen and a basolateral membrane in contact
with the blood supply (50). Sigrist-Nelson and Hopfer (51) first demonstrated with brush
border membrane vehicles isolated from rat intestinal mucosa that fructose absorption is
associated with a saturable carrier unrelated to the sodium dependent glucose transporter.
In 1992, Davidson et al. reported that fructose transporter, glucose transporter 5 (GLUT
5) (41), is expressed at the brush border membrane of the human intestine. Burant et al.
(1992) measured GLUT5 specificity, and reported that GLUT5 exhibits selectively for D-
fructose transport, not D-glucose or D-galactose, with a Km of 6-15 mM in Xenopus
oocytes (52, 53). GLUT5 is not sodium-dependent so it is not inhibited by phlorizin, an inhibitor of sodium dependent glucose transporter (52). In addition, fructose uptake at the brush border membrane by GLUT5 was not inhibited by cytochalasin B, a competitive inhibitor of facilitative glucose transporter (GLUT2) in the human intestine (53). To determine whether GLUT5 prefers fructose in either a pyran or a furan ring form, fructose transport rate by GLUT5 was measured in the presence of fructose analogues
(53). Both L-sorbitol and 1, 5-anhydromannitol (pyran form) did not inhibit fructose uptake by GLUT5. However, 2, 5-anhydromannitol (furan ring fructose analogue) decreased fructose uptake to approximately 30%, indicating that GLUT5 prefers fructose in the furan ring form (53).
In addition to the presence of GLUT5 in the brush border membrane of the small intestine, another fructose transporter has been discovered that can transport fructose across the basolateral membrane of the enterocyte. This transporter, GLUT2, mediates
23 not only the transport of fructose (Km = 16 mM), but also glucose (Km = 30 mM) and galactose (54, 55). GLUT2, like GLUT5, is sodium independent. GLUT2 is inhibited by cytochalasin B (52) and phloretin (55).
Figure 2.2. Fructose absorption in the small intestine (53)
A couple of studies were conducted in rats to measure the intestinal absorption of fructose (56, 57). In vivo, rats were then placed in airtight glass chambers, and different doses of fructose were administered by gastric gavage. The hydrogen amount of the air excreted from each rat was analyzed by gas chromatography. The rats readily absorbed
0.4 g (1.4-1.6 g/kg body weight) of fructose. However, malabsorption was observed with
0.6 g (2.1-2.4 g/kg body weight) of or more of fructose (56).
Studies on fructose absorption in human intestine in vivo are scarce, and they have been conducted via either perfusion study (58) or breath hydrogen tests (59-62).
24 The results of studies assessing fructose absorption via breath hydrogen testing will be
reviewed first and then mechanistic studies of fructose absorption will be addressed.
Clinical assessment of fructose absorption: breath hydrogen testing
Breath hydrogen tests have been used to detect the malabsorption of various
carbohydrates, most often lactose and fructose. The underlying principle of the breath
hydrogen test is that bacterial fermentation of carbohydrates represents the only source of
molecular hydrogen (H2) in the human body. Upon malabsorption of a carbohydrate
from the small intestine, bacteria (predominantly in the large intestine) ferment the
carbohydrate, producing gases such as hydrogen (and sometimes methane) as well as
other products (e.g., short-chain fatty acids). Hydrogen is not metabolized by humans
and variable portions of the hydrogen are excreted in breath and flatus (63). Thus, a rise in breath hydrogen above the fasting level (typically a 10-20 ppm increase) indicates that
carbohydrate has reached the colon (64)
There are several conditions that must be satisfied to accurately assess fructose
absorption by breath hydrogen testing. First, hydrogen-producing bacteria must be
present in the colon. Lactulose, a nonabsorbable disaccharide of galactose and fructose,
can be used to identify the presence of bacteria producing significant amount of hydrogen
(63). Using a sensitive analytical equipment and accounting for possible bacterial
conversion of hydrogen to methane (via the co-measurement of hydrogen and methane),
it is rare to identify a person who does not produce hydrogen in response to malabsorbed
carbohydrate (64).
Second, the foods eaten the day before the breath test may false the fasting breath
hydrogen level. Slowly digested foods, such as beans and high fiber cereals, should not
25 be eaten as a dinner the day before the breath test because they can result in colonic hydrogen production separate from the hydrogen produced by the carbohydrate of interest. In general, white rice and meat are recommended as a dinner to be eaten on the night before the test because both are easily digested, contributing no residual hydrogen that might complicate the interpretation of the test (65). Appropriate fasting (minimum of 10 h before the test) helps to assure a low baseline breath hydrogen value as well.
Third, exercise and smoking may provoke hyperventilation, hence they must be discontinued before or during a test. Previous studies showed that cigarette smoking caused massive increase (3- to137-fold increase) in breath hydrogen concentrations because cigarette smoke contains hydrogen gas (66, 67). Sleeping during the test will increase breath hydrogen levels due to the slowing of intestinal motility, so carbohydrate movement throughout the colon also becomes slow, permitting a longer time for hydrogen accumulation (68). In addition to decreased intestinal motility, sleeping causes reduced rate of air turnover in the lungs, which slows down the rate of hydrogen removal from the blood.
Finally, antibiotics can alter the results of breath hydrogen testing in various ways.
In some cases, broad-spectrum antibiotics can eliminate hydrogen producing bacteria (68,
69), but in other cases antibiotics such as Neomycin can increase breath hydrogen excretion, possibly due to the killing of hydrogen consuming bacteria in the colon (68).
A measurement of carbon dioxide in the breath sample can be used to ascertain the degree of contamination of the breath sample with atmospheric air during the collection process. Because atmospheric air contains virtually no carbon dioxide and the partial pressure of carbon dioxide in alveolar air is 40 torr (5.3 kPa, or roughly 5.5%), the
26 carbon dioxide content of the breath sample is a reflection of how much mixing of the sample has occurred with atmospheric air (which also contains no significant hydrogen or methane concentrations). The hydrogen content of breath samples can be adjusted for variations in the carbon dioxide (CO2) content of the breath sample using the following formula: [(5.5 / % CO2 in the sample) × the measured H2 value of the sample](69).
The breath hydrogen test has been widely used to indicate carbohydrate malabsorption because it is noninvasive and convenient. However, the breath hydrogen test is only a semi-quantitative indicator of the amount of carbohydrate reaching the colon (63). A procedure that is sometimes used for estimating the amount of carbohydrate malabsorbed is to: 1) quantify the amount of breath hydrogen excretion from a particular carbohydrate over a given time frame; 2) compare that amount with the breath hydrogen excretion from an equal dose of lactulose administered over the same time frame. This approach may not be valid due to differences in the rate of fermentation of different fibers, but it is often used because the only other alternatives are invasive intubation techniques or studies in ileostomists.
Fructose absorption capacity in humans
Fructose absorption capacity has been determined as measured via breath hydrogen testing in healthy humans. When fructose dose was increased from 25 g to 50 g in 10% solution in healthy adults, the prevalence of fructose malabsorption increased from 0% to 37.5% (40), 50% to 80% (61), and 19% to 58% (59). Similarly, healthy children aged between 1 month and 17 years after fructose intake at 2 g/kg body weight experienced incomplete absorption of fructose, and children’s age was not significantly correlated with the degree of fructose malabsorption (60). Not only fructose dose but also
27 fructose concentration affected fructose malabsorption. When fructose concentration was
doubled from 10% to 20%, the frequency of fructose malabsorption was also almost
doubled (from 37.5% to 71.4%) (40). These studies suggest that intestinal fructose
absorption capacity is limited, is dose- and concentration-dependent, and is not dependent
on age.
The fructose absorption capacity in humans, when expressed relative to body
weight, is lower than in rats. Fructose amount ranging from 0.5 to1.0 g/kg body weight in
humans caused malabsorption (40, 59, 61), whereas at least 2.0 g fructose/kg body
weight in rats was necessary to induce fructose malabsorption (56). Therefore, care must
be taken in extrapolating results from animal studies (mostly in rats) to humans (61).
Dietary factors affecting fructose absorption
Glucose
Numerous animal and human studies have conducted to determine if absorption
of fructose is improved in the presence of glucose (40, 56-58, 61). The absorptive
capacity of fructose in the presence of either glucose or galactose was measured in rats
(56). Administration of 0.8 g fructose to rats by gastric gavage caused a large amount of
hydrogen excretion, but the feeding of 0.8 g of either glucose or galactose along with same amount of fructose produced significantly lower hydrogen excretion compared with the feeding of fructose alone. The facilitating effect of glucose on fructose absorption was dose-dependent, with the highest fructose absorption capacity obtained with the simultaneous intake of equimolar amounts of fructose and glucose (56). The increased absorption capacity for fructose in the presence of glucose was also reported in perfused isolated rat intestine (57).
28 In human studies, fructose intestinal absorption capability was increased when the mixture of equimolar amount of glucose and fructose was given or when the same amount of fructose was provided as sucrose. The results of human studies of improved fructose absorption are presented in Table 2.1.
29 Carbohydrate amount / Percentage of subjects
Volume who had malabsorption
Fructose (59) 50 g/500 mL 58
25 g/250 mL 19
Fructose + Glucose 25 g + 25 g/250 mL 0
Sucrose 50 g/500 mL 0
Fructose (62) 25 g/200 mL 53
50 g/200 mL 73
Fructose (40) 50 g/500 mL 37
Sucrose 50 g/500 mL 0
Fructose (60) 2 g/kg body weight 71
Fructose + Glucose 2 g/kg body weight 14
+ 2 g/kg body weight
Fructose (61) 50 g/500 mL 80
Fructose + Glucose 50 g + 50 g/500 mL 0
50 g + 25 g/500 mL 30
50 g + 12.5 g/500 mL 70
Sucrose 100 g/500 mL 0
Table 2.1. Fructose absorption capability studies in healthy humans
30 Amino acids
Hoekstra et al. (70) found that fructose absorption was enhanced by L-alanine, L- phenylalanine, and L-proline. L-alanine improved fructose absorption better than other amino acids mentioned above and the effect of L-alanine on improvement of fructose absorption was more profound than glucose.
Fructose, sorbitol, and carbohydrate malabsorption
Carbohydrate malabsorption, as indicated by the breath hydrogen test, has been observed in children with chronic nonspecific diarrhea following a single intake of 8 oz apple juice (71). Interestingly, their symptoms were resolved after the removal of apple juice from their diet in that study. Fruit juices contain varying amounts of fructose, glucose, and sorbitol (Table 2.2) (72). Sorbitol is naturally present in certain fruits and fruit juices, such as prune, pear, and apple (72). Sorbitol is not completely absorbed in the small intestine, and appears to interfere with fructose absorption (21, 61). Apple juice contains sorbitol and more than twice as much fructose as glucose (21). The increased breath hydrogen responses and gastrointestinal symptoms following apple juice intake may be due to fructose malabsorption, sorbitol malabsorption, or the interaction of fructose and sorbitol (73). The least breath hydrogen was produced following the intake of white grape juice, consistent with the fact that white grape juice does not contain sorbitol at all and has low fructose:glucose ratio (74). Rumessen and Gudmand-Hoyer
(73) found significantly increased breath hydrogen levels after the feeding of a mixture of fructose and sorbitol in amounts that cause no malabsorption when administered separately in both healthy adults and patients with functional bowel disease (75). It was suggested that the presence of sorbitol interferences with fructose absorption or vice
31 versa (73). Sorbitol could possibly interact with GLUT 5, reducing fructose absorption or perhaps GLUT 5 is needed for sorbitol absorption and fructose occupies GLUT 5 in
place of sorbitol (73). However, the potential interaction of fructose with other sugar
alcohols has not been described.
Fructose Glucose Sorbitol
Apple juice 6.2 2.7 0.5
White grape juice 7.5 7.1 0.0
Table 2.2. Carbohydrate composition of fruit juices (g/100 mL serving) (72)
Facilitation of fructose absorption by glucose: proposed mechanisms
Disaccharidase-related transport system
The facilitating effect of glucose on fructose absorption has been observed in the
presence of phlorizin. Phlorizin, an inhibitor of sodium dependent glucose transporter
(SGLT1), had no impact on glucose-stimulated fructose absorption at a concentration of phlorizin of either 1 or 2 mmol/L, despite the fact that glucose transport was reduced at 1 mmol/L phlorizin (57). This indicated that a Na+ dependent glucose transporter is not
associated with the facilitating effect of glucose on fructose absorption. The finding that
the addition of the same amount of 3-O-methyl glucose, a glucose derivative that is
transported by the Na+ dependent glucose transporter, to fructose solution did not
32 enhance fructose absorption in rats also supports that a Na+ dependent glucose transporter
is not involved in the glucose-induced facilitation of fructose absorption (56).
Acarbose, an inhibitor of intestinal α-glucosidase including sucrase-isomaltase, significantly inhibited absorption of both sucrose and the mixture of fructose and glucose in rats (56) and in perfused isolated rat intestine (57). Also, sucrose competitively
inhibited glucose-induced increase of fructose absorption when sucrose was added to a
perfusion solution containing both fructose and glucose, suggesting that sucrose
competes with fructose and glucose for the same mechanism of absorption (56, 57). In
these studies, the absorption rate of fructose from the mixture of equimolar amounts of
fructose and glucose was similar to the absorption rate of fructose from a corresponding
amount of sucrose (56). Therefore, Fujisawa et al. (56) and Rumessen and Gudman-
Hoyer (61) proposed the hypothesis that when fructose and glucose are consumed
simultaneously, they may be transported by the disaccharidase-related transport system
on the brush border membrane of the small intestine as if they were the products of the
digestion of sucrose. In addition, it was suggested that the disaccharidase-related
transport system might play an important role in facilitated fructose absorption in the
presence of either glucose or galactose and that the presence of glucose might be crucial
for enhanced fructose absorption in rats.
In contrast with results in the studies conducted by Fujisawa et al. (56), the
inhibition of sucrase-isomaltase activity by acarbose did not impede glucose-facilitated
fructose absorption in adults (58) and children (70). Shi et al. (58), conducting an
intestinal perfusion study with a triple lumen tube in healthy adults, demonstrated that the
presence of free glucose enhanced fructose absorption by 29% compared with no glucose
33
Solvent drag mediated paracellular pathway
Intestinal epithelial cells serve as barriers to the passive transepithelial movement of hydrophilic solutes. Since the permeation of hydrophilic solutes through lipid bilayers is limited, the major pathway of hydrophilic solute transport is paracellular, except in the case of molecules that have intestinal transporters. The epithelial intercellular tight junction is a barrier separating the lumen from the interstitium, thus serving as a rate- limiting barrier of solute transport through a paracellular pathway (50).
34
Figure 2.3. Schematic illustration of the trans-and para-cellular pathway in the epithelial cells. (50)
In 1987, Pappenheimer et al. proposed a new theory of the small intestinal nutrient absorption. They suggested that the presence of sodium-cotransported nutrients in the small intestinal lumen triggers the opening of tight junction for mass transport of water and hydrophilic nutrients by a paracellular pathway (76-78). Pappenheimer et al. demonstrated that the presence of 25 mM glucose in the lumen of rat small intestine increased creatinine absorption by 193% (76). Fluid absorption in the presence of luminal glucose was also twice of that in the absence of glucose (76). Another study showed that the addition of either 25 mM glucose or 12 mM alanine to the perfusion solution caused two- to threefold decreases of transepithelial impedance (77). Decreased
35 transmucosal impedance indicates increases in membrane surface area and width of intercellular junctions (77). Consistently, electron microscopic examination of tight junction structure demonstrated junctional dilatation accompanied by condensation of microfilaments in the zone of the perijunctional actomyosin ring in the presence of either glucose or alanine (78). These data suggest that the presence of active sodium- cotransported nutrients, such as glucose, on the apical membrane of epithelial cells leads to decreased transepithelial resistance (i.e. increased tight junction permeability) via the condensation of the perijunctional cytoskeleton. This increases local osmotic pressure in the lateral intercellular spaces, generating the osmotic force to drive paracellular flow
(79). Turner et al. showed that decreased transepithelial resistance after active Na+- glucose cotransport increased the permeability to mannitol, but not to the larger tracer inulin, meaning that Na+-glucose cotransport-dependent increased permeability is limited
to substances of small molecular weight (80).
One major morphological change in tight junction area before an increase in tight
junction permeability is the condensation of the perijunctional cytoskeleton. A ring of actin and myosin II encircles the apical side of epithelial cells, so the condensation of
perijunctional cytoskeleton, initiated by active Na+-dependent nutrient transport, results in its contraction and increased tight junction permeability, and it has been suggested that there is an association between the cytoskeleton and tight junction permeability (80).
Apical GLUT2
The apical GLUT2 pathway, an alternative view to paracellular pathway, is proposed to explain glucose and fructose absorption within minutes when high concentrations of glucose and/or fructose are present at the apical side of the small
36 intestine (81). When there is little glucose in the lumen in fasting state, glucose
concentration in the lumen is less than 5mmol/L in plasma. After a meal, glucose
concentrations in the lumen range from 50-300 mmol/L (81) within 30 min. Sodium
dependent glucose transporter saturates at 30-50 mM glucose in vitro, but glucose
absorption increases almost linearly to several hundred millimolar (82). Hence, it is likely
that there is another mechanism for the glucose absorption in addition to active Na+- dependent glucose cotransport. Previous kinetic studies reported that glucose is absorbed by two components; one is saturable and phlorizin-sensitive with Km of 17.9 – 22.6 mM and the other pathway is non-saturable and phlorizin-insensitive (83, 84). In the apical
GLUT2 pathway, the latter is considered a GLUT2-mediated passive component.
Before a meal, GLUT2 amount is very low at the brush border membrane (Figure
2.3) and GLUT2 at the basolateral membrane provides glucose from blood to maintain the energy requirement of the enterocytes. After a meal, glucose level at the brush border
membrane is high after hydrolysis of starch and disaccharides. The increased glucose
transport across the apical membrane occurs via SGLT1, resulting in activation of protein
kinase C (PKC) ßII. The activation of PKC ßII leads activation of GLUT2 already
present in the apical membrane and translocation of GLUT2 present at the basolateral
membrane to the apical membrane (81). In addition to glucose transport by SGLT1,
fructose uptake by GLUT 5 also triggers PKC ßII, casuing activation of GLUT2
translocation into the apical membrane (85). Apical GLUT2 is then the major pathway of
glucose and/or fructose absorption. As glucose concentration in the lumen decreases, the
signaling pathway is reversed so that GLUT2 is inactivated and traffics away from the
apical membrane (81). GLUT2 translocation to brush border membrane is associated with
37 SGLT1 activation so that SGLT1 is seen to play a critical regulatory role as well as its
function as a transporter (86).
Figure 2.4. The apical GLUT2 model of glucose absorption (A) before a meal and (B) after a meal (81).
Insertion of GLUT2 from the basolateral membrane to the brush border membrane is associated with activation of protein kinase C (PKC) ßII. In brush border membrane vesicles prepared from rat jejunum, not only GLUT2 levels but also GLUT2 intrinsic activity increased with glucose concentration. PKC ßII level in the brush border
38 membrane also increased with increasing glucose concentration. SGLT1 is another
glucose transporter in the brush border membrane, so it was determined if inhibition of
SGLT1 would suppress levels of PKC ßII and GLUT2. Phloridzin, an inhibitor of SGLT1, was added to perfusion buffer, and it was observed that inhibition of SGLT1 with phloridzin decreased GLUT2 level in the brush border membrane and proportionally diminishes GLUT2-mediated glucose transport (87). Similar results were reported when in perfusion buffer, NaCl was replaced with choline chloride and Na+ in the buffer salts
was replaced with K+; sodium replacement caused a fast first phase of SGLT1 inhibition
and a slow second phase of GLUT2 inhibition with decreased PKC ßII (87).
GLUT2 transports not only glucose but also fructose (88). It was investigated if
fructose absorption in the apical membrane was mediated by GLUT2. Wild type mice
were fed a standard chow diet or a fructose rich (65%) diet for 5 days or were fed with a single gastric bolus of 40% fructose, glucose or mixture of fructose and glucose after overnight fast. After sacrifice, intestinal brush border membrane vehicles were prepared from the mice, and western blots were conducted to measure GLUT2 expression in brush border membrane from the mice after fed different diets. GLUT2 was expressed at the brush border of mice adapted to the high fructose diet and mice fed bolus of 40% fructose, glucose or mixture of fructose and glucose (85). Thus, it seems that fructose also
activates GLUT2 translocation from basolateral membrane to brush border membrane in
mice. To determine if PKC is involved in the fructose-activated GLUT2 insertion to the brush border membrane, an in vitro perfusion of rat small intestine was conducted. The jejunum of rat was perfused for 30 min in vivo and then 60 min in vitro with 5 mM fructose with or without phorbol 12-myristine 13-acetate (PMA, a PKC activator). The
39 perfusion of rat jejunum with fructose and PMA increased fructose absorption to 1.7 fold compared with the absence of PMA. This result was consistent with the level of GLUT2 at the brush border membrane of rat jejunum perfused with 5 mM fructose in the presence of PMA; PMA increased GLUT2 levels to 3.84-fold those of the PMA-absent treatment. This increased GLUT2 level at the brush border membrane was associated with a 4-fold increase in GLUT2-mediated transport. However, GLUT5 levels and
GLUT5-mediated fructose absorption was not changed. Therefore, the increase in
GLUT2-mediated fructose transport is regulated by PKC (88).
Erythritol-Introduction
Erythritol is a tetrose sugar alcohol (1,2,3,4-butanetetrol) (Fig 2.4.) with a sweetness of 60-80% that of sucrose (89). Erythritol is naturally found in some fruits, mushrooms, and fermented foods such as wine, sherry, and soy sauce (see Table 2.3).
Erythritol can also be manufactured from glucose by fermentation with osmophilic yeasts
(Moniliella sp., Trichosporonoides sp.) or fungus (Aureobasidium sp.) (90, 91). The estimated dietary intake of erythritol is 1.3 g/person/day (89).
CH2OH | H---C---OH | H---C---OH | CH2OH
Figure 2.5. Chemical structure of erythritol (MW=122.12 g/mol) (89)
40 Foods Erythritol content
Grapes 0-42 mg/kg
Melons 22-47 mg/kg
Pears 0-40 mg/kg
Miso bean paste 1310 mg/kg
Sake 1550 mg/L
Sherry wine 70 mg/L
Soy sauce 910 mg/L
Wine 130-300 mg/L
Table 2.3. Concentrations of erythritol in various foods (90).
Erythritol can be combined with other sweeteners to reduce the caloric content of products and to improve taste and texture. Erythritol has been used in chewing gum, confectionery, beverages, and sugar substitutes in Japan since 1990 (89). A Generally
Recognized As Safe (GRAS) affirmation petition of erythritol was accepted by the US
Food and Drug Administration on January 1997 (92). Canada also has permitted the use of erythritol as a sweetener in diabetic foods, candies, and chewing gum (93). Recently,
European countries have approved the use of erythritol in foods (94).
41 Erythritol Absorption
Erythritol absorption has been studied in animals and in humans (95-98). Because
absorbed erythritol is not metabolized by the body (89, 99), the urinary erythritol
excretion reflects the absorption rate (96).
Studies in rats have examined the effects of dose and pre-adaptation on the degree
of absorption and excretion of erythritol (100, 101). When Wistar rats were fed on a diet containing 5% erythritol ad libitum for 28 days, approximately 92% of administered erythritol was excreted into the urine and 1% in the feces. When erythritol amount was increased to 10% in the diet, urinary excretion of erythritol was lowered to 83%, but fecal excretion remained same at 1% (100). In an effort to examine the metabolic fate of erythritol precisely, the same authors used 14C-erythritol in rats (101). When rats were fed
14C-erythritol orally by gavage at a dose of 0.1 g/kg body weight, 88% of 14C-erythritol
was excreted intact in the urine within 24 hours, and only 6% of the labeled carbon from
14 erythritol was excreted as CO2 during the same time period, mainly due to erythritol
fermentation by the colonic microflora (101). The proportion of labeled carbon in expired
CO2 within 24 hours was increased to 10% when rats fed the same level of erythritol
were “adapted” to it (per the feeding of 10% erythritol in the diet for 2 weeks) before the
study. It is unclear whether such adaptation will occur when humans consume large
amounts of erythritol for extended periods of time (101). In non-adapted rats, the dose of
14C-erythritol was increased from 0.1 g/kg body weight to 1.0 g/kg body weight. It was observed that urinary erythritol excretion within 24 hours in rats fed the 1.0 g/kg body
weight dose was four times higher than those fed the dose of 0.1 g/kg body weight.
42 Erythritol absorption has also been measured in humans. Renal excretion of
erythritol, an indicator of erythritol absorption in humans, is similar to that observed in
rats. In a study measuring urine kinetics of erythritol in six healthy humans who were administered a single oral dose of erythritol at 1.0 g/kg body weight (average amount of administered erythritol =64 g/person), the cumulative urinary erythritol excretion was 2%,
7%, 20%, 30%, and 78% for 0.5, 1, 2, 3, and 24 hour postprandial times, respectively
(95). When the dose of erythritol was lowered to 0.3 g/kg body weight (average dose was
17.3 g/person), cumulative urinary erythritol excretion within 24 hours was
approximately 88% (98). It is likely that small doses of erythritol are more readily
absorbed than high doses when erythritol is provided as a single oral dose in solution. In
another study, six healthy subjects each consumed a snack containing either 0.4 or 0.8 g
erythritol/kg body weight, and urine was collected for 22 hours postprandial times.
Erythritol appeared in the urine within 2 hours after intake of erythritol-containing snack.
At the end of 22 hours, 61% and 62% of the ingested amount of erythritol was recovered
in the urine of the individuals administered 0.4 and 0.8 g erythritol/kg body weight,
respectively (95). Urinary recovery of erythritol after intake of a snack containing erythritol was lower compared with intake of erythritol in solution. This discrepancy
must be caused by slower gastric emptying of snack (solid food) than a solution.
Although urinary collection period in the latter study in which a snack containing
erythritol was administered was 22 hours, not 24 hours, it may not result in lower urinary
excretion of erythritol because only 1% of erythritol intake/hour during the period from 8 to 24 hours after intake of erythritol solution (98).
43 Erythritol Distribution
The distribution of erythritol in body fluids and tissues has been measured in
animal studies. In Wistar rats after a single administration of 14C-erythritol in doses
ranging from 0.125 to 2.0 g/kg body weight, the maximum plasma erythritol levels were
reached at 1 hour after administration regardless of dose (102). The peak plasma erythritol concentrations increased dose-dependently, except for the highest dose (2.0 g/kg body weight). The peak plasma erythritol value at 2.0 g/kg body weight was approximately only 1.5 times higher than that at 1.0 g/lg body weight, indicating saturation of absorption from the small intestine (102). In rats fed a single oral dose of
14C-erythritol at 1 g/kg body weight, 55% and 15% of the administered radioactivity was
found in the stomach and small intestine , respectively. Radioactivity levels were also
high in the kidney, bladder, and liver, organs that are involved in erythritol absorption
and excretion (102).
The plasma kinetics of erythritol has been examined in healthy humans. Six healthy adults (mean age 32.7 ± 6.8 y, BMI 22.0 ± 1.7 kg/m2 ) were administered a single
aqueous oral dose of erythritol at 1.0 g/kg body weight (average amount of administered
erythritol was 64 g/person). Plasma erythritol levels increased steadily, reaching mean
maximum value of 18.0 mmol/L at 90 min postprandially. The levels then declined
gradually to approximately 12.3-13.9 mmol/L at 3 hours postprandially (97). Plasma
erythritol levels measured by Noda et al. (98) with six subjects after intake of 0.3 g of
erythritol /kg body weight reached the peak value (3.5 ± 1.0 mmol/L) at 30 min
postprandially. In another study, plasma erythritol levels were measured in six healthy
subjects following the administration of a snack containing either 0.4 or 0.8 g
44 erythritol/kg body weight. In the group having 0.4 g erythritol /kg body weight, plasma erythritol level was reached peak at 3.0 ± 0.84 mmol/L at 1 h postprandial (98).
Maximum plasma erythritol concentration in 0.8 g/kg body weight was 5.1 ± 0.7 mmol/L,
but occurred at 2 hours after intake of erythritol-containing snack. Compared with the
first two studies, peak plasma erythritol levels appeared to be lower following administration of erythritol in snack than in aqueous solution. It may be caused by slower gastric emptying related with solid type of snack compared with solution (103). Also, the small number of subjects (n=6) in each study may not be enough to reflect true
differences in plasma erythritol levels between different treatments.
Metabolism of malabsorbed erythritol by the colonic microflora
In an in vitro 6-h fermentation of 14C-erythritol with cecal contents from rats fed
10% erythritol diet for 2 weeks, approximately 60% of added 14C-erythritol was
fermented to CO2 and short chain fatty acids including acetic acids, propionic acids, and
butyric acids (101). Approximately 72.7%, 67%, and 72.6% of administered erythritol
was recovered in the urine 24 hours after administration of 0.1 g/kg body weight 14C- erythritol in germ-free, conventional pre-adapted and conventional non pre-adapted rats,
14 respectively (96). CO2 excretion in feces was very small in germfree rats (0.8 %), but it
was substantial in conventional adapted (10.9%) and in conventional non-adapted rats
14 (6.7%). The higher expiration of CO2 in adapted rats than non -adapted rats can be explained by adaptation of the colonic microflora to erythritol. It also means that
14 erythritol is not easily utilized by nonadapted microflora. The considerable rates of CO2 excretion in conventional adapted rats indicate that ingested erythritol is fermented by colonic microflora to absorbable and metabolizable intermediates because colonic
45 adaptation can improve the ability to ferment malabsorbed erythritol. However,even after adaptation, only 10 % of fed 14C-erythritol was excreted, so it is probably likely that a very high amount of erythritol is required for colonic adaptation (96).
The metabolism of erythritol was measured in healthy subjects after intake of 25 g
13 13 C-erythritol (104). No CO2 and H2 excretion in breath samples collected over 8 h
(indices of colonic bacterial metabolism) were detected, indicating that erythritol was not used as a substrate of colonic bacterial fermentation. A high proportion (84%) of the administered dose of erythritol was recovered in urine after 24 hours, indicating that only a small proportion of the ingested erythritol might have become available for colonic
13 fermentation. However, the lack of excretion of either CO2 or H2 in the breath suggests that the erythritol was not fermented. An in vitro study was undertaken to confirm that erythritol is nonfermetable. A dose of 50 mg of erythritol was incubated with fresh fecal flora under anaerobic conditions for 6 hours and H2 production was measured. Consistent
13 with the vivo study, no H2 and CO2 were produced by fecal flora from erythritol (105).
Another study was performed to investigate if erythritol is fermented when an incubation period is prolonged to 24 hours. For the study, erythritol was incubated with human inoculate for 24 hours and not only erythritol disappearance but gas and short chain fatty acids production was measured. Neither gas nor short chain fatty acids were produced during in vitro fermentation of erythritol. The result that erythritol is not fermentable is consistent with when the fermentation period was prolonged to 24 hours (105).
46 Effect of erythritol on glycemic and insulinemia
Serum glucose and insulin were not affected by administration of 0.3 to 1 g
erythritol/kg body weight in healthy adults (97, 98). In addition, serum glucose and insulin levels were remained unchanged after intake of 20 g erythritol in five patients
with type 2 diabetes (106). Given the finding of no response of serum glucose and insulin
for erythritol, the GI and insulin index values of erythritol are both 0 (103).
Laxative effect of erythritol
In a study to determine the laxative threshold of erythritol in humans, 66%
subjects experienced diarrhea with the intake of 62.5 g erythritol as a jelly (107).
However, a single ingestion of 25 g erythritol as a jelly did not cause diarrhea in humans
(107). The laxative threshold of erythritol in female subjects was 0.8 g/kg body weight,
and 0.66 g/kg body weight for males. 50% female subjects complained nausea and 23%
experienced flatus after ingestion of 0.85-1.2 g erythritol/kg body weight. In this study,
subjects consumed erythritol in a jelly form, and it was questionable if the laxative
threshold of erythritol may be different between jelly and liquid forms. There was no
significant difference observed in the fecal condition between jelly and solution intake
containing 50 g erythritol (107). A recent study reported that subjects consuming 20 g
and 35 g erythritol in a liquid did not experience any gastrointestinal symptoms, such as
nausea, bloating, borborygmi. However, when erythritol consumption was increased to
50 g, a significant increase in nausea and borborygmi was observed (108).
Urinary excretion of Na, K, and Cl is not affected by 0.3 g/kg body weight
erythritol intake in humans (98), which indicates little osmotic diuretic effect of 0.3 g/kg
body weight erythritol. No adverse gastrointestinal symptoms were reported at the doses
47 in the study (98). However, osmotic diuretic effect of erythritol was reported with 4 g/kg body weight/day in rats (100).
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60 CHAPTER 3
DETERMINATION OF THE GLYCEMIC AND INSULIN INDEX VALUES
OF RAISINS IN THREE POPULATIONS
Introduction
The glycemic index (GI) is a system of ranking foods according to how much
they raise blood glucose relative to a reference food (1). The insulin index is determined in a similar manner, except that blood insulin concentrations are used instead of blood
glucose levels (2). A number of studies have reported that high postprandial blood insulin levels in response to a high GI food are related with an increase in obesity (3-6),
cardiovascular disease (5-7), and diabetes (5,6, 8, 9). Insulin increases cholesterol
synthesis in the liver via increased activity of HMG-CoA reductase, and inhibits the
activity of hormone-sensitive lipase, thus enhancing lipogenesis that are related with the
progression of obesity and other chronic disease (10). Hence, it appears that the
knowledge of the GI of a food may be augmented by further data on the postprandial
insulin response.
Low GI foods are recommended as a pre-exercise snack to athletes because they
provide stable and prolonged blood glucose and insulin levels during exercise, possibly
resulting in improved performance (11). In addition, acarbose, which lowers GI of foods
61 by inhibiting digestion of carbohydrate, reduced the progression rate from prediabetes to
diabetes (12).
Raisins are nutritious snack, containing dietary fiber, antioxidants, potassium, and
iron (13). Raisins are also a concentrated source of carbohydrate (a 2 T., or 18 g, portion
counts as a fruit exchange in the Exchange Lists for Meal Planning)(14). As such, there
could be potential concerns about raisins causing a high postprandial glycemic response,
especially in persons with diabetes or prediabetes. However, both the glycemic index (GI)
of a carbohydrate source and the absolute amount of carbohydrate in the food portion
influence the postprandial glycemic response (1). Even though raisins are a concentrated source of carbohydrate, roughly half of their available carbohydrate is fructose (15),
which has a GI value of just 19 (glucose = 100) (16). The other half of carbohydrate in
raisins is glucose (15). A GI value for raisins of 64 ± 11 (glucose = 100) in healthy adults
was reported by Jenkins et al. (1) and another study reported a GI value of 65.7 ± 5.8
(glucose = 100) in gestational diabetes women (17). Some important criticisms of these 2
studies, however, include a small sample size of 6 subjects in the former study, the use of
non-standard blood sampling times in the latter study, and no measurement of the insulin response in either study.
A key tenet of GI methodology is that factors such as the presence of diabetes or
differences in the exercise habits and physical conditioning of the subjects will not affect
the GI value. This is because the glycemic response to the test food is compared with the glycemic response to a reference food (glucose solution or white bread) within the same subject in a GI study (18,19). However, recent investigations have suggested that the GI
of some carbohydrate sources may be substantially lower when measured in athletically
62 trained versus untrained persons (20, 21). Therefore, the first objective of this study was
to measure the glycemic and insulin indexes of raisins according to standardized
methodology. The second objective was to determine if the glycemic and insulin index
of raisins differ when measured in 3 groups of people who are in different metabolic
status : 1) a group of sedentary young and healthy adults; 2) a group of young aerobically
trained adults; and 3) a group of adults with prediabetes.
Methods
Subjects
The study groups comprised 10 healthy sedentary individuals (S), 11 aerobically
trained young adults (A), and 11 prediabetic adults (P). The mean age (± standard error of
the mean) of the S group was 25.7 ± 1.3 years, with a body mass index (BMI) of 23.3 ±
1.7 kg/m2. The mean fasting plasma glucose level of the S group was 87.2 ± 1.7 mg/dL.
The mean age of the A group was 23.1 ± 1.0 years. This group had a normal mean BMI
(24.1 ± 0.3 kg/m2) and a normal fasting plasma glucose level (87.6 ± 2.3 mg/dL). The
mean age of the P group was 50.0 ± 2.6 years, with a mean BMI of 32.6 ± 1.9 kg/m2, and a mean fasting plasma glucose level of 110.5 ± 2.6 mg/dL. Groups S and A reported no history of glucose intolerance, diabetes, gastrointestinal disorders or recent use of antibiotics. Subjects in group S reported that they had not exercised more than 3 hours per week for the last 6 months. The subjects in the A group reported, by virtue of their responses to a questionnaire (22) that they had trained aerobically more than 8 hours per week for the past 6 months. Three subjects in the A group were rowers, seven were swimmers, and one was a runner. The subjects in the P group had a history of glucose
63 intolerance (fasting plasma glucose level between 100 and 125 mg/dL) (23), and they had
exercised less than 3 hours per week for the past 6 months.
The study protocol was reviewed and approved by the Institutional Review Board
Human Subjects Committees at the Ohio State University (Columbus, OH), and the State
University of New York at Brockport (Brockport, NY). All subjects provided written
informed consent for the study.
Feeding Protocol
The study was a two-treatment, randomized, crossover study with a minimum of
3 days between each treatment visit. Subjects were asked to consume at least 150 g per
day of carbohydrate for 3 days prior to each visit and refrained from vigorous exercise for
12 hours prior to each visit. At each visit, subjects arrived at the laboratory after having
fasted overnight at least 10 hours. A baseline capillary blood sample was collected via a
finger puncture in the fasting state. The subjects then consumed either 296 mL of a
glucose tolerance test beverage containing 50 g glucose (SunDex®, Fisher Health Care,
Houston, TX) or a 69-g portion of raisins (Sun-Maid®, Kingsburg, CA) that was
calculated to provide 50 g of available carbohydrate (total carbohydrate minus dietary fiber) based on the nutrition label information. The order of the test meals was randomized. The energy and macronutrient composition of the 50-g available carbohydrate-portion of raisins was as follows: Energy 224 kcal, 0 g fat, 1.7 g protein,
53.5 g carbohydrate, dietary fiber 3.5 g, 50 g sugars. The glucose solution provided 200
kcal (50 g carbohydrate as glucose, 0 g protein, 0 g fat).
64 Collection and analysis of serum glucose and insulin
Finger-stick capillary blood samples were collected using sterile lancets at
baseline (immediately before ingestion of the raisins or glucose solution), and at 15, 30,
45, 60, 90, and 120 min (and 150 and 180 min for the P group) postprandially. Timing
started at the first bite/sip of the raisins/glucose solution (24). Approximately 1 mL of
whole blood was obtained from each finger-prick and collected into serum separator
tubes (Becton Dickinson, Franklin Lakes, NJ). Blood was allowed to clot and then
centrifuged at 1168 x g for 15 min to obtain serum. Serum glucose concentrations were
analyzed using the YSI 2700 Select Plus Biochemistry Analyzer (Yellow Springs
Instruments, Yellow Springs, OH) via the glucose oxidase method. Serum insulin was
analyzed by enzyme-linked immunosorbent assay (ELISA) using an Insulin DSL-10-
1600 ACTIVE kit (Diagnostic Systems Laboratories, Inc., Webster, TX).
Calculations of area under the curve
The positive incremental area under the curve (IAUC) for serum glucose and
insulin was calculated geometrically. Any area beneath the fasting values was ignored
(24). Excursion for serum glucose and insulin (difference between each individual’s
maximum and minimum serum glucose and insulin values), and baseline-adjusted peak
(BAP) glucose and insulin (difference between each individual’s peak and baseline serum glucose and insulin values) were also calculated for each group.
65 Statistical analysis
Descriptive statistics were calculated and normality tests were performed for all
variables using the NCSS 2000 software package (NCSS Computing, Kayesville, UT).
Data are displayed as the mean ± SEM. Data that were non-normally distributed were transformed (square root or logarithmic) prior to statistical analysis to approximate a normal distribution. ANOVA for a randomized block design was used to determine global significant differences. In the event of a significant ANOVA result (P < 0.05), the
Tukey-Kramer post-hoc test was used for pairwise comparisons (25).
Results
The serum glucose and insulin responses to the test foods in groups S, A, and P
(left to right) are shown in Figure 3.1. There were no significant differences in baseline
serum glucose levels for the both the glucose solution and raisin meals among the three
groups. The serum glucose response to the raisins was significantly lower than for the
glucose solution at several postprandial time points in all three groups. Although the
average glucose responses to the raisins were virtually identical in the S and A groups,
the glucose solution resulted in a serum glucose curve that was higher relative to the
raisins at 90 min in the S group versus the A group.
The serum glucose IAUC for both the raisins and the glucose solution was not
significantly different among the 3 groups. However, the serum glucose IAUC of the
glucose solution was 14% lower for the S group and 31% lower for the A group
compared with the P group (Table 3.1). The GI of raisins was low (GI ≤ 55) in the S
66 (49.4 ± 7.4) and P (49.6 ± 4.8) groups and was moderate (GI 55-69) in the A group (62.3
± 10.5), but there were no differences among the groups (P = 0.437).
The serum glucose excursion in response to glucose solution in the P group was
higher than the S and A groups (P < 0.001 each). Although there were no significant
differences in the GI of raisins based on glucose excursion, the GI of raisins based on
glucose excursion in the P group was 24% lower for the S group and 19% lower for the A
group. The BAP value for both raisins and glucose solution was not significantly
different among the 3 groups. The GI of raisins based on BAP in the P group was 15%
lower for the groups of S and A groups. In the S group, both glucose excursion and BAP
resulted in higher GI values as compared to IAUC. In addition, the P group had a
significantly higher GI based on BAP when compared with the GI based on IAUC.
However, there was no difference in the GI of raisins based on different calculations in
the A group.
The insulin responses to both the glucose solution and the raisins followed the
same trends as the glucose responses in each of the three groups, with the raisins resulting
in significantly lower insulin values at several time points compared to glucose. The peak serum glucose and insulin responses for both the raisins and the glucose solution were higher in the P group compared with groups S and A. The A group had lower
serum insulin AUC for both glucose solution (P = 0.002) and raisins (P = 0.008) than P
group (Table 3.2).
Serum insulin AUC for the glucose solution in S group was 2.2 times that of the
A group, but it was not significantly different (P = 0.075). A similar trend was observed
for serum insulin response to raisins in groups S and A, and the response in S group was
67 90% higher than group A. Insulin index of raisins was not significantly different among
groups (P=0.72). The insulin index of raisins were 47.3 ± 9.4, 51.9 ± 6.5, and 54.4 ± 8.9
for S, A, and P groups, respectively. However, the A group secreted 2-2.5-fold less
insulin per gram of carbohydrate compared with the S and P groups, respectively
(P<0.05).
The A group had lower serum insulin IAUC in response to both glucose solution
(P = 0.002) and raisins (P = 0.008) compared with P group. These data corresponded with
the excursion values (P = 0.012 and P = 0.004, respectively) and the BAP values (P =
0.01 and P = 0.008, respectively). Serum insulin IAUC in response to glucose solution in
the S group was 2.2 times that of the A group, but it was not significantly different. Both
insulin excursion and BAP in the S group were 2 times that of the A group. The insulin
index based on excursion was very similar with that based on BAP in all 3 groups. The
insulin index of raisins based on IAUC tended to be lower than the insulin index based on
excursion and BAP.
Discussion
The GI of raisins did not vary significantly among the different populations in our
study, although the GI of raisins in athletes tended to be higher than in sedentary people
and prediabetic subjects. Because the GI is a ratio of the IAUC values of the test and
reference foods, the GI could be elevated by either a high IAUC for the test food or a
lower than expected IAUC for the reference food. Because the A group had the lowest
serum glucose IAUC for the raisins among the three groups, the somewhat higher GI of
raisins in the A group was associated with a less pronounced increase in serum glucose
68 following the glucose solution. When the GI of raisins was calculated using excursion and BAP, the GI of raisins in people with pre-diabetes appeared to be higher than the other groups. It must be due to significantly higher glucose excursion in response to glucose solution in the P group compared with the other groups, whereas glucose excursion and BAP in response to raisins were similar among the groups.
Glucose and insulin excursion (the difference between maximum and minimum) and BAP (the difference between maximum and baseline) were used as alternative ways to calculate GI and insulin index of raisins in the present study. Numerous studies have demonstrated the importance of glucose excursion and BAP in association with the risk of micro-macrovascular disease through mechanisms of oxidative stress and endothelial
dysfunction (26-28). The Diabetes Control and Complications Trial Research Group recommended not only the monitoring of hemoglobin A1c, but also the reduction of acute blood glucose fluctuations (i.e., postprandial glycemia) to prevent and manage
micro- and macrovascular complications in type 2 diabetes (29). A limitation in the use
of IAUC to calculate postprandial glycemia/insulinemia is that, as a single number
summary, it does not provide information on the shape of the blood glucose curve. The
IAUC may not appropriately distinguish between a food that causes a rapid spike and
subsequent quick decline in glucose from a food that causes a more moderate and
sustained elevation .. Thus, the use of IAUC with excursion/BAP may provide more
accurate information to monitor and control postprandial glycemia/insulinemia.
It is likely that increased insulin sensitivity in athletes results in faster glucose
disposal from a large glucose challenge than for sedentary people or prediabetic people
(30-33). In our study, athletes had 47-53% less insulin secretion for both the raisins and
69 the glucose solution compared with sedentary and prediabetic people. Enhanced insulin sensitivity in athletes is mediated not only by increased GLUT4 translocation to the skeletal muscle cell membrane but also by increasing GLUT4 levels, thus increasing the glucose uptake into the muscle (34-36) . Two different signal pathways to trigger
translocation of GLUT4 in the muscle have been suggested. The insulin-dependent
pathway involves increases in insulin-stimulated insulin receptor substrate-1 (IRS-1)
tyrosine phosphorylation and phosphatidylinositol 3-kinase (PI 3-kinase) (34, 35). The
other pathway to increase glucose uptake in the muscle is insulin-independent, but
contraction-dependent. Muscle contraction during exercise increases the AMP/ATP and
creatine/creatine phosphate ratios, resulting in an increase in AMP-activated protein
kinase (AMPK) activity (36). The increased AMPK activity is associated with
contraction-stimulated, increased glucose uptake in muscle. However, the signaling
pathway for how AMPK activation may be related with GLUT4 translocation is not
known.
In contrast, insulin resistance is commonly found in prediabetes, type 2 diabetes,
and obesity. Insulin resistance results from decreases in tyrosine kinase activity, IRS-1 tyrosine phosphorylation, and PI 3-kinase activation. GLUT4 protein content in adipose
tissue is reduced in obesity and type 2 diabetes (35). However, GLUT4 protein content is
normal in muscle from the people with type 2 diabetes and obesity, although GLUT4 translocation is reduced (35). It is likely that the pre-diabetic subjects in this study
suffered from one or more of these defects in insulin signaling. In addition to the effects
of lifestyle on insulin sensitivity, age is also negatively associated with insulin sensitivity
(37) and our prediabetic subjects were significantly older than the other subjects.
70 However, a number of case-control studies reported that the increased insulin resistance
with age is caused by age-related changes in body composition (i.e. increase in body fat
and decrease in lean body mass) and the decrease in physical activity, rather than by age per se (37, 38). Insulin resistance in the people with impaired fasting glucose resulted in postprandial hyperglycemia by a decrease in insulin-stimulated glucose uptake by peripheral tissues.
Previously, GI values of raisins were reported as 64 ± 11 (glucose =100) in 6 healthy
subjects (1) and 65.7 ± 5.8 (glucose =100) in 6 women with gestational diabetes (17) .
These GI values of raisins are comparable with the GI based on IAUC in athletes (62.3 ±
10.5) and they are higher than the GI of raisins in groups of S and P in this study.
However, only 6 subjects participated in each of the previous studies. In addition, raisins
were provided with tea made with milk in the first study (1). The exact composition of
the tea and milk were not specified and there may have been factors in the tea that altered
the GI measurement. In the study in the women with gestational diabetes (17), the GI of
raisins was calculated using less than the standard of 7 blood samples, and the blood
samples were collected for 2 h postprandially, which was not long enough for the blood
glucose level dropped down around the baseline. It may affect the GI of raisins in the
previous studies.
Two recent studies showed that the GI of breakfast cereals was dependent on the training status of subjects, with a lower GI of cereals reported in trained subjects (20, 21).
However, in our study, the GI of raisins was not significantly different among groups,
although there was a somewhat higher GI value in the athletically trained subjects. The
reason(s) for the discrepancy between our results and the results of Mettler et al. (20, 21)
71 is/are are not clear. One variation between the studies is that the cereals in their studies were fed with partially skimmed milk bringing the protein content to 10-14 g and the fat content to 5-9 g. By contrast, the meals in our study were nearly 100% carbohydrate.
Currently, it is unknown if macronutrients could have differential effects on glycemia in trained versus untrained persons. However, there is some evidence that dietary fiber and protein may lower glycemia more in persons with high versus low waist circumference, and that fat reduces glycemia more in those with low versus high fasting plasma insulin
(39).
One important difference between our results and those of Mettler et al. (20) is the higher glycemic AUC for the glucose solution in our population of sedentary adults
(271 mmol/min/L) versus theirs (208 mmol/min/L). This is in contrast to the relatively similar glycemic AUC values for athletes in our study (217 mmol/min/L) compared with theirs (202 mmol/min/L) for the glucose solution. Some of the discrepancy in the sedentary values between these studies may be due to their reporting of whole blood glucose values versus serum glucose in our study. Whole blood glucose concentrations are typically 10-15% lower than plasma or serum values (40). However, even accounting for this difference, there may have been important subject factors that influenced the differential GI results. It should be noted that in their second study of cereals (21), the mean whole blood glucose AUC in response to the same 50-g glucose solution as the first study (with the same glucose monitoring system) was ~ 175 mmol/min/L. Pooling our results with those of Mettler et al. (20, 21), it is apparent that even when studying healthy sedentary subjects of similar mean age (23-26 y) and BMI (21-23 kg/m2), it is possible
72 for the mean glycemic AUC to a 50-g glucose solution to vary from ~175 to 271
mmol/min/L.
One of the limitations of this study was small subject number. In this study, 10-11
subjects per group participated to measure the glycemic and insulin index of raisins.
Although FAO/WHO recommends having at least 10 subjects per group in a GI test, insufficient sample size may be responsible for lack of statistical significance of results
and high values of standard errors. Use of more subjects would have resulted in higher
statistical power and less variance between subjects within the group.
In conclusion, this study demonstrated that the GI of raisins is low to moderate,
despite their concentrated carbohydrate content. In addition, the GI of raisins was not
significantly different when measured in healthy sedentary individuals, athletes, and
prediabetic persons. Our findings contradict other recent studies, indicating that further
research is needed to evaluate the effect of training status on GI. Finally, our study
confirms that endurance-trained athletes are able to normalize postprandial glycemia with
lower insulin secretion compared with healthy sedentary adults.
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78 Group S Group A Group P 12 * 12 * * 12 11 11 11 10 Glucose 10 * 10 * Raisins * * 9 9 * 9 * 8 8 * 8 * 7 7 * 7 6 6 6 Serum glucose(mmol/L) 5 Serum glucose (mmol/L) 5 5 Serum glucose (mmol/L) 4 4 4
0 Time30 postprandial60 (min)90 120 Time postprandial (min) 0 30Time 60 postprandial 90 (min) 120 150 180 0306090120 79 * * * 90 90 * 90 80 Glucose 80 80 * 70 Raisins 70 70 60 60 60 * 50 50 ** 50 40 40 * * 40 30 30 * 30 20 Serum insulin (uIU/mL) insulin Serum 20 20 Serum insulin (uIU/mL) insulin Serum (uIU/mL) insulin Serum 10 10 10
0 0 0
Time postprandial (min) Time postprandial (min) Time postprandial (min) 03060901200 30 60 90 120 0 30 60 90 120 150 180
Figure 3.1. Serum glucose and insulin responses to raisins and glucose solution in sedentary (Group S), athletically trained (Group A), and pre-diabetic (Group P) subjects. Data points are the mean ± SEM.
* indicates significant differences between glucose solution and raisins ( P < 0.05).
Group S Group A Group P
Glucose Glucose Glucose Raisins solution Raisins solution Raisins solution IAUC1 125.0 ± 19.6 270.5 ± 33.0 113.3 ± 15.1 217.3 ± 30.5 148.5 ± 23.7 314.2 ± 54.3 (mmol/min/L)
GIIAUC 49.4 ± 7.4 --- 62.3 ± 10.5 --- 49.6 ± 4.8 ---
2 2 2 80 Excursion (mmol/L) 3.2 ±0.3 4.5 ±0.5 3.1 ±0.3 4.6 ±0.4 4.0 ±0.3 6.9 ±0.5
GIExc 77.0 ±6.5 --- 71.8 ±6.2 --- 58.6 ±6.5 ---
BAP (mmol/L) 2.9 ±0.2 4.1 ± 0.5 2.8 ±0.2 4.0 ± 0.4 3.1 ±0.2 5.1 ± 0.5
GIBAP 76.1 ±7.6 --- 76.3 ±7.2 --- 64.3 ±7.6 ---
Table 3.1. Positive incremental areas under the curve (IAUC), excursion, and baseline-adjusted peak (BAP) for serum glucose and corresponding glycemic index of raisins in the S group (N=10), the A group (N=11) and the P group (N=10) when calculated using each type of data. Data represent the mean ± SEM.
1Raisin IAUC< glucose solution IAUC for Group S ( P = 0.001), Group A ( P = 0.006), and Group P ( P = 0.012)
2Raisin excursion significantly different between A and P groups ( P < 0.001)
Group S Group A Group P
Glucose Glucose Glucose Raisins solution Raisins solution Raisins solution IAUC 1 1938 ± 399 4625 ± 1143 1021 ± 125 2 2068 ± 217 3 2431 ± 414 2 5110 ± 851 3 (μIU/min/L)
Insulin Index IAUC 47.3 ± 9.4 --- 51.9 ± 6.5 --- 54.4 ± 8.9 ---
Excursion ( μIU/L) 36.3 ±5.0 65.6 ±10.5 21.8 ±4.3 4 33.5 ±8.9 5 44.5 ±4.5 4 72.9 ±9.4 5 Insulin index Excursion 62.4 ±10.2 --- 68.8 ±8.7 --- 67.6 ±9.2 --- 81 BAP ( μIU/L) 36.2 ±5.1 65.3 ± 10.4 21.4 ±4.4 6 32.8 ± 8.8 7 42.3 ±4.5 6 70.0 ± 9.3 7
Insulin index BAP 62.1 ±10.5 --- 69.3 ±8.9 --- 67.3 ±9.4 ---
Table 3.2. Positive incremental areas under the curve (IAUC), excursion, and baseline-adjusted peak (BAP) for serum insulin and corresponding insulin index of raisins in the S group (N=10), the A group (N=11) and the P group (N=10). Data represent the mean ± SEM. 1Rasin IAUC < glucose solution IAUC for Group S ( P = 0.029), Group A ( P < 0.001), and Group P ( P = 0.014) 2Raisin IAUC significantly different between A and P groups ( P = 0.008) 3Glucose solution IAUC significantly different between A and P groups ( P = 0.002) 4Raisin excursion significantly different between A and P groups ( P = 0.004) 5Glucose solution excursion significantly different between A and P groups ( P = 0.012) 6Raisin excursion significantly different between A and P groups ( P = 0.008) 7Glucose solution excursion significantly different between A and P groups ( P = 0.01)
CHAPTER 4
INHIBITION OF FRUCTOSE ABSORPTIONI BY ERYTHRITOL
IN HEALTHY ADULTS
Introduction
Fructose may be a beneficial sweetener for people with diabetes because of its
low glycemic index (fructose’s glycemic index = 19, where glucose = 100) (1). Further,
several studies have shown that ingestion of 7.5-10 g of fructose with or before a
carbohydrate challenge (e.g., glucose solution or mashed potato meal) reduces the
glycemic response to the carbohydrate challenge (2-4). It is also known, however, that
fructose, when fed alone, is often poorly absorbed from the gastrointestinal tract (5-7).
This can lead to gastrointestinal distress that may limit the potential usefulness of fructose as a sweetener.
Presently, there is incomplete knowledge regarding intestinal fructose absorption
in humans. A highly specific intestinal transporter for fructose (GLUT5) has been identified (8, 9), but it is possible that other pathways of intestinal fructose transport may
be involved. It is well-known that fructose absorption increases dramatically with the co-
ingestion of an equimolar amount of glucose (representative of the products of sucrose
hydrolysis). In rats, fructose absorption appears to be coupled with glucose via a
82 disaccharidase-related transport system (10). However, Shi et al. (11) reported that such
a system does not exist in humans and suggested that the facilitation of fructose
absorption by glucose occurred via the paracellular route.
Erythritol is a 4-carbon sugar alcohol that is essentially noncaloric (0.2 kcal/g),
non-glycemic, and very well absorbed (>90%) from the intestine (12). No intestinal
transporter for erythritol has been identified and it is presumably absorbed via the paracellular route (perhaps some degree of transcellular passive transport as well).
Erythritol is used in pharmaceutical studies to evaluate absorption of drugs and there is one report of the ability of erythritol to increase the paracellular absorption of calcium
(13). Given the desirable nutritional properties of erythritol (noncaloric, nonglycemic)
and its presumed paracelullar absorption, we hypothesized that erythritol might make a
desirable facilitator of fructose absorption via the paracellular pathway suggested by Shi et al. (11). The primary objective of this study was to explore if simultaneous ingestion
of erythritol with fructose increases fructose absorption in healthy subjects as measured
by breath hydrogen concentrations. The secondary objectives were to: 1) determine the
postprandial serum fructose responses to beverages containing 50 g of fructose with or
without erythritol; 2) measure the postprandial blood glucose responses to beverages
containing 50 g of fructose with or without an equimolar amount (33.3 g) of erythritol;
and 3) determine total rectal gas passages, flatus, and subjective gastrointestinal tolerance
in the postprandial period to beverages containing 50 g of fructose with or without an
equimolar amount of erythritol.
83 Methods
Subjects
Sixty-two healthy subjects from The Ohio State University (Columbus, OH)
community were enrolled, and 37 subjects (13 men and 24 women) completed the study.
Of the 25 subjects who exited from the study prematurely, 2 subjects became ineligible
for the study as they required antibiotic therapy, 13 subjects withdrew due to inability to
tolerate blood draws, and 10 were unable to continue participation due to their busy
schedules. The subjects who completed the study are described as follows: mean age (±
standard error of the mean) of 23.0 ± 0.5 years, mean body mass index of 22.8 ± 0.4
kg/m2, and mean fasting plasma glucose level of 4.8 ± 0.1 mmol/L. All subjects were
healthy, free from infectious, metabolic, and gastrointestinal diseases, and did not take
any antibiotics for 3 weeks prior to the study. Subjects also did not take any medications
or dietary supplements at doses known to affect glycemia or gastric motility. In addition,
all subjects produced breath hydrogen (>20 ppm rise above baseline) in response to 10 g
lactulose ingestion (14). This ensured that hydrogen-producing bacteria were present in
these individuals (15, 16). The self-reported ethnicity of the subjects who completed the study was: 27 Caucasians (73%), 7 Asian or Pacific Islanders (19%), 2 African
Americans (5%), and 1 Hispanic (3%). The study was approved by the Western
Institutional Review Board (Olympus, WA) and informed consent was obtained from all subjects before start of the study.
Feeding Protocol
The study was a randomized, double-masked, crossover design. Subjects
participated in three separate 3-hour beverage tolerance tests. They were instructed to
84 consume at least 150 g of carbohydrate per day for 3 days before each visit, and refrain
from exercise the day before each visit. A low-residue standard dinner was provided to
them between 4 and 7 pm on the day before each visit. The standard low-residue dinner consisted of 240 mL of Ensure Plus® along with variable quantities of Ensure® Nutrition and Energy Bars (Abbott Laboratories Ross Products Division, Columbus, OH) such that the products provided a total caloric value equal to one third of each subject’s estimated daily energy requirement (based on the Harris-Benedict equation multiplied by a light activity factor of 1.3) (17). At each visit, subjects consumed one of the three test beverages consisting of 500 mL of water plus the following sugars, administered in random order: 50 g fructose (F), 50 g fructose and 50 g glucose (FG), and 50 g fructose and 33.3 g erythritol (FE). Subjects consumed the study product within 10 min. Subjects were allowed to drink up to 240 mL of water during the 3-h beverage tolerance test.
Data collection and analysis
Blood collection
For blood collection, one of the subject’s hands were placed in a
thermostatically-controlled hand box in which the air temperature was maintained
precisely at 50º C to obtain arterialized blood (18). The hand was kept in the hand box for
the entire duration of the meal tolerance test. An indwelling catheter was inserted into a
dorsal hand vein, and arterialized blood was collected from the heated hand vein at
baseline, and at 15, 30, 45, 60, 90, 120, 150, and 180 minutes after administration of the
study product. Whole blood samples were allowed to clot and then centrifuged at 1168 ×
g for 15 min to obtain serum.
85 Analysis of serum glucose, lactate, insulin, fructose and erythritol
Serum glucose concentrations were analyzed using the YSI 2700 Select Plus
Biochemistry Analyzer (Yellow Springs Instruments, Yellow Springs, OH) via the
glucose oxidase method. Serum insulin levels were determined by enzyme-linked
immunosorbent assay (ELISA) using an Insulin DSL-10-1600 ACTIVE kit (Diagnostic
Systems Laboratories, Inc., Webster, TX). Serum fructose and erythritol concentrations
were measured by a high-pH anion exchange chromatograph with pulsed amperometric
detection (HPAEC-PAD) using a Dionex 500 system (Dionex, Sunnyvale, CA) at Ross
Products Division Abbott Laboratories (Columbus, OH).
Breath hydrogen collection and analysis
Breath samples were collected at baseline and hourly for 8 h after administration
of the study product. Subjects were instructed to collect their breath samples in sealed
evacuated tubes (Exetainer, Labco International, Inc., Houston, TX) using an
AlveoSampler mouthpiece (Quintron Instrument Company, Milwaukee, WI). Breath
samples were analyzed for hydrogen, methane, and carbon dioxide concentrations by gas
chromatography (Quintron Microanalyzer Model SC, Quintron Instrument Company,
Milwaukee, WI.). The observed hydrogen and methane values were corrected for atmospheric contamination of alveolar air by normalizing the concentrations of observed carbon dioxide to 5.26% (5.3 kPa), which is the partial pressure of carbon dioxide in alveolar air (19). Subjects were identified as having carbohydrate malabsorption if their breath hydrogen concentrations increased more than 20 ppm for 8 h compared with basal
nadir value (the lowest breath hydrogen value at baseline, 1 or 2 h postprandial) (14).
86 Rectal gas passage, bowel movement and stool consistency
Subjects recorded the total number of rectal gas passages (flatus) using a
counting device (VWR International, West Chester, PA) for 0 through 8 hours after
administration of study product. The number of bowel movements for 24 hours after
administration of study product was also recorded. The stool consistency for each bowel movement was rated using a 5-point scale: 1 = hard/dry-pellets, small, hard mass; 2 = hard/formed-dry, stool remains form and soft; 3 = soft/formed-moist, softer stool that
retains shape; 4 = soft/unformed-stool pudding like; 5 = watery-liquid that can be poured
(20).
Gastrointestinal symptoms
Subjects recorded the frequency and intensity of gastrointestinal symptoms
(nausea, abdominal cramping, distension and flatulence) for 0 through 24 hours after the
study product consumption using 10-cm visual analog rating scales (0 = usual or absent,
10 = more than usual or severe). This scale has been used to measure subjective GI
symptoms in previous studies (21, 22).
Calculations of area under the curve
The positive incremental area under the curve (AUC) was calculated
geometrically for serum glucose, fructose, insulin, lactate, erythritol, and breath hydrogen
concentrations. Any area beneath the fasting values was ignored (23).
87 Statistical analysis
A power analysis, using data from a previous study (5), was conducted which indicated that 36 subjects would be required to detect a 47 % difference in the incremental AUC for breath hydrogen with 84% power. Data are presented at the mean ±
SEM. Data, after examination for normality, were analyzed using ANOVA for a randomized block design to test for global significant differences. If the global ANOVA was significant, the Tukey-Kramer post-hoc test was used for pairwise comparisons (24).
Results were considered statistically significant only if the P-value of an analysis was <
0.05.
Results
Breath hydrogen
The changes in breath hydrogen concentration are shown in Figure 4.1. No significant differences were found among beverages for the baseline breath hydrogen values. The FE beverage caused significantly higher breath hydrogen levels at 1 h (P <
0.001), 2 h (P < 0.001), 3 h (P < 0.001), and 5 h (P = 0.021) postprandial compared with the FG beverage. Lower breath hydrogen values at 1 h (P < 0.001) and 2 h (P < 0.001) postprandial were observed with the FG beverage compared with the F beverage. In addition, positive incremental breath hydrogen AUC was significantly different across treatments (P < 0.001) (Table 4.1). The AUC for the FE beverage was twice the AUC of the F beverage and nine times the AUC of the FG beverage.
88 Serum fructose
There were no significant differences in baseline serum fructose levels among the beverages (Figure 4.2). The FE beverage resulted in lower serum fructose concentrations at 30 min (P < 0.001), 45 min (P < 0.001), and 60 min (P = 0.009) postprandially compared with the F beverage. Serum fructose levels were significantly higher with the FE beverage at 45 min (P = 0.024) and lower at 120 min (P < 0.001) versus the FG beverage. Serum fructose levels were significantly higher for the F beverage at 30 min (P = 0.005), 45 min (P < 0.001), and 60 min (P < 0.001) postprandial compared with the FG beverage (). The positive incremental serum fructose AUC was higher for the F beverage compared with the other two beverages (P < 0.001 in each case).
Serum erythritol
The baseline serum erythritol levels were not significantly different among beverages (Figure 4.3). The FE beverage increased the serum erythritol level from near zero to approximately 5 mmol/L within 60 min and this level was significantly higher compared with non-erythritol beverages (FG & F) (P < 0.001). No significant differences in serum erythritol were found between the FG and F beverages.
Serum glucose
There were no significant differences among beverages in the serum glucose levels at baseline (Figure 4.4). However, serum glucose levels at 15 min (P < 0.001), 30 min (P < 0.001), 45 min (P < 0.001), 60 min (P < 0.001), 90 min (P < 0.001), and 120
89 min (P < 0.001) postprandial were significantly higher with the FG versus beverages FE
and F. There were no significant differences in serum glucose levels between the FE and
F beverages. Positive incremental serum glucose AUC was significantly higher than FE
and F (P <0.001) (Table 4.1). The positive incremental serum glucose AUC was lowered
by 90 % for the FE (P < 0.001) and by 82% for the F (P < 0.001) beverages when
compared with the FG beverage.
Serum insulin
The insulin response to the treatments is illustrated in Figure 4.5. There were no significant differences at baseline for serum insulin among the beverages. Serum insulin concentrations were increased in subjects consuming the FG beverage compared with the other treatments at 15, 30, 45, 60, 90, 120, and 150 min postprandial. There was a higher insulin response at 60 min with F compared with FE (P = 0.039). When the AUC values of the FG group were compared with the F & FE groups, a trend similar to that observed with serum glucose AUC was evident (Table 4.1). Both the F and FE beverages had serum insulin AUCs that were 82% lower than the AUC for FG beverage.
Serum lactate
The baseline serum lactate levels were not significantly different among
beverages (Figure 4.6). The FE beverage resulted in significantly lower serum lactate levels at 30 min (P < 0.001), 45 min (P < 0.001), 60 min (P < 0.001), 90 min (P < 0.001),
and 120 min (P < 0.001) postprandially compared with the FG beverage. No significant differences in serum lactate levels were found between FE and F beverages except at 30
90 min (P = 0.012). Serum lactate levels were significantly higher with the FG beverage at
30 min (P < 0.001), 45 min (P = 0.002), 90 min (P = 0.005), and 120 min (P = 0.003)
postprandially compared with the F beverage. Serum lactate AUC of the FG beverage was significantly higher than the F and FE beverages (Table 4.1).
Gastrointestinal Symptoms
The number of rectal gas passages for 8 hours after ingestion of study product was
significantly greater (P < 0.001) for FE group compared with FG group (Table 4.2). For
the FE beverage, the total number of rectal gas passages was 22% higher compared with
F group, but this increase was not significant (P = 0.128). Compared with FG and F, the
FE beverage resulted in significantly higher numbers of bowel movements (P < 0.001 for each comparison). In addition, the FE beverage caused more watery stool compared with the other beverages (P < 0.001 for each comparison).
Discussion
There were both expected and unexpected findings in this study. As expected, the
feeding of 50 g fructose in the absence of additional glucose resulted in poor fructose
absorption. This finding is in agreement with several studies in the literature (5-7). Also
as expected, the combination of equimolar amounts of fructose and glucose greatly
enhanced fructose absorption. This is reflected in both the lack of a significant breath
hydrogen response to the FG beverage and also in the improved gastrointestinal tolerance
of the FG versus the F beverage. Several studies in the literature have produced similar
findings (5-7). The key disadvantage to the facilitation of fructose absorption by glucose
91 is the large glycemic and insulin responses that were observed. Such a large glycemic
response would not be desirable in persons with diabetes. It should be noted that
although the FG beverage markedly showed lower serum fructose level than the F
beverage, this finding is not due to fructose malabsorption (given the lack of breath
hydrogen response). Serum lactate, a major product of fructose metabolism, AUC was
significantly higher with the FG group compared with the F and FE groups, meaning that
fructose is well absorbed with the FG group compared with the F and FE group. Instead,
it is likely that the large insulin response may have driven fructose uptake by extra-
hepatic cells. GLUT5 is known to be expressed in insulin-sensitive tissues such as
skeletal muscle or adipose (25), but it is unclear if GLUT5 itself is directly upregulated
by insulin.
At present, it is not fully understood how the presence of glucose can facilitate the
intestinal absorption of fructose. One potential explanation for the enhancing effect of
glucose on fructose absorption is that the presence of glucose or other active sodium-
cotransported nutrients on the apical membrane of epithelial cells leads to decreased
transepithelial resistance (i.e. increased tight junction permeability) via the condensation
of the perijunctional cytoskeleton, generating osmotic force to drive paracellular flow
(26). This hypothesis is supported by the data of Shi et al. (11) that were discussed earlier.
Another possible explanation for the facilitating effect of glucose on fructose
absorption is that increased luminal glucose concentrations could promote the
translocation of GLUT2 within the enterocyte. Normally, GLUT2 is present mainly on
the basolateral side of the enterocyte and can transport both glucose and fructose to the
92 blood. However, under conditions of high luminal glucose availability, GLUT2 may translocate to the apical membrane and assist in the transport of either glucose or fructose
(27).
Clearly, the most unexpected finding was that erythritol in the FE beverage did not enhance fructose absorption and, in fact, likely impaired it. Perhaps the increased osmolality of the FE beverage relative to the F beverage might have decreased small intestinal transit time, resulting in looser stools and increased gas production. Subjects reported increased stool frequency and more liquid stools in the 24-h period after the FE beverage compared with the F or FG beverages. However, it is difficult to accept that increased osmolality is the main explanation for the results. If increased osmolality/rapid intestinal transit was the primary explanation, it would be expected that the absorption of both fructose and erythritol would be decreased. However, the serum erythritol level rose significantly in the FE beverage from baseline, reaching a peak of approximately 5 mmol/L within 60 min postprandially. This serum erythritol level is comparable, when adjusted for dose, with a previous study in which volunteers that were fed 0.3 g/kg (mean of 17.4 g/person) erythritol (about half the size of the dose fed in our study) reached a maximum plasma erythritol level of 2.2 mmol/L by 90 min post-ingestion (28). One limitation of our study was the lack of an erythritol-only beverage, which would have allowed a more complete evaluation of serum erythritol levels. If erythritol were absorbed 100% in the FE beverage, the maximum serum erythritol level would have been approximately 7-9 mmol/L. In previous studies measuring the plasma kinetics of erythritol, mean maximum plasma erythritol was 18 mmol/L and 3.5 mmol/L after average intake of erythritol of 64 g/ person and 17.3 g erythritol /person, respectively.
93 Rumessen and Gudmand-Høyer (29) observed that combinations of fructose and
sorbitol increased carbohydrate malabsorption in 7/10 subjects who did not previously
experience malabsorption with either carbohydrate fed individually. No differences in
small intestinal transit time were observed for the fructose-sorbitol mixtures. As in our
study, the breath hydrogen technique employed in their study can not distinguish between
fructose and a sugar alcohol (sorbitol or erythritol) as the identity of the malabsorbed
carbohydrate. However, in our study, we have strong evidence that it was primarily fructose, not erythritol, that was malabsorbed. The FE beverage caused a large increase in serum erythritol as well as decreases in serum fructose and lactate compared with the F beverage. In addition, several in vitro fermentation studies with human fecal samples
demonstrated no increase in hydrogen product with the addition of erythritol, indicating
that it is poorly fermented by colonic bacteria (30, 31). Finally, it was reported in a
recent study that 35 g of erythritol in a liquid (almost same dose fed in our study) did not
cause any gastrointestinal symptoms or waterly stools (32).
An intriguing hypothesis for our findings is that erythritol and fructose may
compete for the same intestinal carrier. A similar hypothesis was proposed for the
fructose-sorbitol interaction described by Rumessen and Gudmand-Høyer (29). Although
it has long been assumed that erythritol is absorbed passively from the gut (mainly via the paracellular route), a recent study (33) has identified a sodium-dependent intestinal
carrier for glycerol, which is another small molecular weight compound that was
previously thought to be absorbed passively. In addition, a new sodium-dependent
glucose transporter (SGLT4) mRNA has been found to be expressed in the human
intestine (34). There is some evidence that SGLT4 can transport either fructose or a
94 sugar alcohol (1,5-anhydroglucitol) in a monkey COS-7 kidney cell line (34). Among the potential candidates for a common carrier of both fructose and erythritol are GLUT5 (the fructose transporter), GLUT2 (which may have a role in fructose uptake when carbohydrate concentrations are high in the intestinal lumen), and SGLT4 (as described above). Further studies using intestinal perfusion techniques or studies in cell culture models and/or brush border membrane vesicles are needed to fully understand the absorption of sugar alcohols in the presence and absence of other carbohydrates.
The interest in the use of erythritol in beverages and confectionery applications has been growing, as there is an increasing demand for reduced-calorie, low-glycemic index products. However, not much is known about the interaction of erythritol and other food components. Fructose is a natural constituent of fruits and is commonly used as an additive in confectionery and soft drinks, possibly in combination with erythritol. The results of this study suggest that food manufacturers need to be aware of potential interactions between sugars and sugar alcohols that can cause gastrointestinal distress in consumers and, therefore, limit product acceptability.
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99 140 a
120
100 a a FG 80 FE F
60 a a
40 b Breath hydrogen (ppm) a
20 b b b b, b 0 012345678 Time postprandial (h)
Figure 4.1. Postprandial breath hydrogen concentrations for all beverages at each time point. FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and
F = 50 g fructose. Values represent mean ± SEM (N = 32-37), and values with different superscript letters at each time point are significantly different (P < 0.05).
100 0.7 a a a 0.6
b b FG 0.5 FE b F
0.4 c ba b
0.3 b
0.2 b Serum fructose(mmol/L)
0.1
0 0 30 60 90 120 150 180 Time postprandial (min)
Figure 4.2. Postprandial serum fructose concentrations for all beverages at each time point. FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and
F = 50 g fructose. Values represent mean ± SEM (N = 32-37), and values with different superscript letters at each time point are significantly different (P<0.05).
101 6
5
4
3 FG FE 2 F Serum erythritol (mmol/L) erythritol Serum
1
0 0 30 60 90 120 150 180 Time postprandial (min)
Figure 4.3. Postprandial serum erythritol concentrations at each time point. FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and F = 50 g fructose.
Values represent the mean ± SEM (N = 32-37). Serum erythritol levels were negligible (<
0.006 mmol/L) for the F and FG beverages, and the FE beverage caused significantly higher plasma erythritol values than the other beverages at all postprandial time points (P
< 0.05).
102
9 a a
a 8 FG FE F a 7
a
6
b b b Serum glucose (mmol/L) b 5 b b b b b b 4 0 30 60 90 120 150 180 Time postprandial (min)
Figure 4.4. Postprandial serum glucose concentrations for all beverages at each time point. FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and
F = 50 g fructose. Values represent the mean ± SEM (N = 32-37), and values with different superscript letters at each time point are significantly different (P<0.05).
103
55
a a a a
45
FG FE a F 35
a
25 a b Serum insulin(uIU/mL)
15 b b b b b
bb c b bb,b b 5 0 30 60 90 120 150 180 Time postprandial (min)
Figure 4.5. Postprandial serum insulin concentrations for all beverages at each time point.
FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and F = 50 g fructose. Values represent the mean ± SEM (N = 32-37), and values with different superscript letters at each time point are significantly different (P<0.05).
104
a 5 a a
ab a b
4 b a FG b b FE b F b 3 b
c b
2 Serum lactate (mmol/L) Serum
1
0 0 306090120150180 Time postprandial (min)
Figure 4.6. Postprandial serum lactate concentrations for all beverages at each time point.
FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and F = 50 g fructose. Values represent the mean ± SEM (N = 32-37), and values with different superscript letters at each time point are significantly different (P<0.05).
105
FG FE F
Breath hydrogen (ppm·h) 1098 ± 269a* 9611 ± 1516b 4636 ± 809c
Serum fructose (mmol·min·L-1) 53.8 ± 1.7 a 48.9 ± 2.1 a 61.0 ± 2.2 b
Serum erythritol (mmol·min·L-1) 0.3 ± 1.0 a 727.7 ± 22.2 b 0.3 ± 1.0 a
Serum glucose (mmol·min·L-1) 304 ± 33 a 30.6 ± 7.3 b 56.0 ± 11.4 b
Serum insulin (µIU·min·L-1) 4306 ± 453a 723 ± 108b 781 ± 114 b
Serum lactate (mmol·min·L-1) 316 ± 21 a 194 ± 16b 220 ± 14b
Table 4.1. Postprandial breath hydrogen, serum fructose, erythritol, glucose, insulin, and lactate areas under the curve (AUC) in response to all study beverages. FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and F = 50 g fructose.
* Values represent the mean ± SEM (N = 32-37), and values with different superscript letters within the same row are significantly different at P < 0.05.
106
No. of rectal gas No. of bowel Average stool consistency
passages per 8 h movements per 24 h ratings **
FG 3.78 ± 0.49a* 1.14 ± 0.14a 2.92 ± 0.17a
FE 10.14 ± 1.48b 2.84 ± 0.40b 4.03 ± 0.16b
F 7.89 ± 1.45b 1.22 ± 0.18a 3.03 ± 0.18a
Table 4.2. Self-reported frequencies of rectal gas passages and bowel movements and stool consistency ratings in the postprandial period in response to each study beverage.
FG = 50 g fructose and 50 g glucose, FE = 50 g fructose and 33.3 g erythritol, and F =
50 g fructose.
*Values represent mean ± SEM (N = 32-37), and values with different superscript letters within the same column are significantly different (P < 0.05).
**5-point stool consistency rating scale, 1 = hard vs 5 = watery
107 CHAPTER 5
ERYTHRITOL INHIBITS TRANSEPITHELIAL TRANSPORT OF FRUCTOSE
ACROSS CACO-2 HUMAN INTESTNAL CELLS
Introduction
Erythritol is a four-carbon sugar alcohol that is essentially non-caloric (0.2 kcal/g)
and non-glycemic (1). However, unlike most other sugar alcohols, erythritol is well
absorbed from the small intestine and is rapidly excreted unchanged in the urine (1).
Several reports suggest a possibility of carrier-mediated transport of erythritol in the
small intestine (2-4). In isolated loops of the small intestine of rats, approximately 30% of erythritol treated was absorbed via the transcellular pathway (2). Fordtran et al. measured water movement and osmolarity of erythritol compared with mannitol, a known
unabsorbable substance, in humans using triple-lumen technique (3). Erythritol exerted
46% less osmotic pressure than its theoretical osmotic pressure across the membrane,
possibly due to carrier-mediated transport of erythritol. In addition, more water
movement was caused by mannitol compared to erythritol. In the jejunum, erythritol was
approximately 60% as effective as mannitol for inducing bulk water flow (3). Finally,
108 carrier (hexose transfer system)-mediated erythritol in red blood cells has been reported
(5).
Because erythritol is also presumed to be absorbed via the paracellular route, our
research group hypothesized that erythritol might enhance paracellular transport of
fructose by opening tight gap junctions (1). Enhanced fructose absorption would be beneficial to prevent diarrhea associated with fructose ingestion (6,7) and may enhance the ability of “catalytic” doses of fructose to lower glycemia (8).
We investigated the effect of erythritol on fructose absorption when an equimolar
amount of erythritol was administered simultaneously with fructose to healthy humans.
Healthy adults (n = 37) consumed one of the following three solutions on three separate
occasions: 1) 50 g fructose and 50 g glucose; 2) a mixture of 50 g fructose and 33.3 g
erythritol, and 3) 50 g fructose. Unexpectedly, the fructose-erythritol treatment resulted in
the greatest increase in breath hydrogen, indicating the most carbohydrate malabsorption.
However, the identity of the malabosorbed carbohydrate, whether fructose or erythritol,
in the fructose-erythritol treatment could not be determined via the breath hydrogen
technique that was employed. In vitro studies (9, 10) have demonstrated that erythritol
was not fermentable by colonic microflora so that it is likely that fructose was the source
of colonic hydrogen production. Further evidence that fructose but not erythritol was
malabsorbed included a considerable rise in serum erythritol and a lowering of the serum
fructose level in the fructose-erythritol treatment compared with the other treatments.
Due to this apparent competition between fructose and erythritol for intestinal
absorption, we hypothesized that erythritol may be absorbed via a transcellular pathway
in the same way that fructose is with the two possibly sharing the same intestinal carrier
109 (11). Thus, experiments are proposed to investigate the effect of erythritol on fructose transport at the cellular level using 3H-fructose and 14C-erythritol.
Caco-2 cells, a human colon cancer cell line, are a widely used model for the
study of the uptake, metabolism and transport of nutrients and drugs because they display
many of the the morphological and functional characteristics of enterocytes (12). They
are polarized with distinct apical and basolateral membranes and express the glucose
transporters SGLT1 and GLUT2, as well as the fructose transporter GLUT5 (12-15).
Fructose uptake and metabolism has been characterized in this cell model (16), and the
effect of erythritol on glucose flux also has been investigated in this cell line (6). The
aims of this study were 1) to determine if erythritol inhibits fructose trasnport in Caco-2 cells and 2) to explore whether inhibition of fructose absorption by erythritol is dose- dependent.
Methods
Materials
The Caco-2 cells were purchased from the ATCC. Dulbecco’s modified Eagle’s
medium (DMEM), Hank’s balanced salt solution (HBSS), amino acid supplements, antibiotics, and fetal bovine serum were purchased from Sigma Chemical Co. (St. Louis,
MO). D-[5-3H]-fructose (specific activity 5 mCi/mmol) and [1-14C]-erythritol (specific
activity 55 mCi/mmol) were obtained from American Radiolabeled Chemicals, Inc. (St.
Louis, MO). Tissue culture flasks (75 cm2) were purchased from Falcon/Becton-
Dickinson (Lincoln Park, NJ), and multiwell dishes and inserts were obtained from Fisher
Scientific.
110
Cell culture
Caco-2 cells were grown and maintained in the laboratory of Dr. Failla. Caco-2
cells from passages 25 through 40 were used all experiments. In brief, stock cultures were
maintained in 75 cm2 flasks in a humidified incubator with an atmosphere of 95% air and
5% CO2 at 37°C to 60-80% confluency. For experiments, detached cells were added to inserts (4.2 cm2, 0.4 µm pores), to 6-well culture dishes at 3.0 x 105 cells/well, and grown
in the presence of DMEM containing 15% FBS, 1% glutamine, 1% NEAA, 1%
penicillin-streptomycin and 0.5 μg/mL amphotericin B. After 40 days, the amount of FBS
was decreased to 7.5 %. Spent medium was removed and fresh medium was added every
other day. Studies were conducted at 17-20 days after seeding cultures. Monolayers were
washed twice with phosphate buffered saline (PBS) at 37°C before adding test media.
Glucose-free, serum-free, high phenol red DMEM supplemented with 3H-fructose and 14C-erythritol insert Caco-2 cells
Phenol red –free, serum-free DMEM containing 5 mM glucose
Figure 5.1. Experiment design. 111 Pilot study
A pilot study was conducted to determine the incubation time of Caco-2 cells treated with fructose for which fructose transport from the apical to the basolateral compartment is the highest. In this study, 2 mL of 5 mM fructose plus 1 µCi of D-5-3H- fructose was added to glucose-free, serum free, high phenol red (500 μM) DMEM in the apical compartment. The basolateral compartment received 2 mL phenol red-free, serum- free DMEM containing 5 mM glucose. Caco-2 cells were incubated for 10, 20, 30, and
60 min in a humidified incubator with an atmosphere of 95% air and 5% CO2 at 37°C
following addition of 2 mL of indicated medium to each chamber.
Experiment 1
This experiment was conducted to determine if fructose transport from the apical
to the basolateral compartment was inhibited by erythritol at high doses (25 mM and 50
mM) of fructose. To initiate the experiment, 2 mL of indicated concentrations of fructose
plus 2 µCi of D-5-3H-fructose with or without 25 mM erythritol plus 1 µCi of 1-14C-
erythritol added to glucose-free, serum free, high phenol red (500 μM) DMEM was added
to the apical compartment. The basolateral compartment received 2 mL phenol red-free,
serum-free DMEM containing 5 mM glucose.
Experiment 2
Experiment 2 was performed to measure if the inhibitory effect of erythritol on
fructose transport was dose-dependent. The integrity of the monolayer was tested one day
prior to the experiment by measuring diffusion of phenol red from the apical to
112 basolateral compartment during a 30 min incubation. The result of phenol red assay close
to 0 %/hr/cm2 indicates the presence of the intact cell monolayer (17). Phenol red flux
was 0.052 %/hr/cm2, indicating barrier integrity of the monolayers. To initiate the
experiment, 2 mL of glucose-free, serum-free, high phenol red DMEM supplemented
with 25 mM fructose plus 2 µCi of D-5-3H-fructose and various concentrations of
erythritol (0-75 mM) plus 1 µCi of 1-14C-erythritol was added to the apical chamber. The
basolateral chamber received serum-free, phenol red-free DMEM containing 5 mM
glucose.
Experiment 3
The purpose of the experiment 3 was to determine if erythritol inhibited fructose
transport following an overnight fast of the Caco-2 cells. The day before the experiment,
2 mL glucose-free, phenol red-free DMEM was added to the apical compartment, and 2
mL of phenol red-free DMEM supplemented with 5 mM glucose and 7.5 % serum was
added to the basolateral compartment. The cells were incubated overnight. The cell
cultures were washed with PBS before the treatments, and glucose-free DMEM with 500
μM phenol red supplemented with 25 mM fructose plus 2 µCi of D-5-3H-fructose with
different concentrations of erythritol plus 1 µCi of 1-14C-erythritol was added to the apical compartment. The basolateral medium contained 2 mL glucose-free and phenol red
-free DMEM.
113 Experiment 4
This experiment was conducted to determine if fructose transport from the apical to the basolateral compartment was affected by erythritol at 1 mM fructose. The apical
medium contained low concentrations of fructose (1 mM) and erythritol (ranging from 0
to 20 mM) supplemented with glucose-free DMEM with high phenol red. Phenol red-free
DMEM supplemented with 5 mM glucose was used for the basolateral side.
For each experiment, cultures were incubated for 30 min in a humidified
incubator with an atmosphere of 95% air and 5% CO2 at 37°C following addition of 2 mL of indicated medium to each chamber. Uptake was terminated by washing twice with
ice-cold PBS. Aliquots (10 μL) from the apical side and 200 μL distilled water were transferred into 5 mL scintillation cocktail (SciniVerse, Fisher Scientific) to determine 3H and 14C. Aliquots (250 μL) from basolateral side were transferred into 5 mL scintillation
cocktail to determine 3H and 14C. Aliquots (300 μL) from basolateral sides were also used
to quantity phenol red. Cells were collected by scraping the insert with a rubber
policeman to re-suspend cells in 1 mL PBS. The insert was washed with another 0.5 mL
PBS and scraped. The suspended cells were centrifuged at 2000 g at 4ºC for 5 min. The
PBS was discarded and cell pellets were sonicated in 1 mL PBS, and stored at 4 ºC for a
maximum of 3 days. Aliquots (100 μL) from sonicated cell pellet were solubulized to 5
mL scintillation cocktail. To measure 3H and 14C, aliquots were dark adapted overnight,
and analyzed by liquid scintillation spectrometry (Beckman LS Model 3801).
114 Statistical analysis
Each experiment was performed with n=3, and all data are expressed as means ±
SEM. Means were compared using one way analysis of variance (ANOVA) for statistical
significance. Results were considered statistically significant when P < 0.05. NCSS 2000 software package (NCSS Computing, Kayesville, UT) was used for statistical analysis.
Results
Pilot study
Fructose transport from the apical to the basolateral compartment increased
proportionally from 10 to 30 min (P=0.000102), but remained unchanged after 30 min incubation (Figure 5.2). Thus, 30 min was chosen as the incubation period for the subsequent experiments.
Experiment 1
After 30 min incubation, 1.68% and 1.34 % of the fructose were transported from
the apical to the basolateral compartments when 25 mM and 50 mM fructose were added
to the apical compartment, respectively (Figure 5.3). The presence of 25 mM erythritol
in the test medium decreased fructose transport in cultures incubated with 25 mM and 50
mM fructose by 24 ± 1.1% and 14 ± 0.7%, respectively. Thus fructose transport was
decreased more with the presence of equimolar amount of erythritol. In contrast,
erythritol flux from the apical to the basolateral compartment was 0.27% of the erythritol
present in the medium containing 25 mM fructose, and this was not altered by the
addition of 50 mM fructose to the medium. Caco-2 cells retained 14.5 nmol and 27.8
115 nmol fructose when incubated with 25 mM and 50 mM fructose, respectively,
corresponding to 0.056% of the fructose in the medium, each. The addition of erythritol
to the apical compartment lowered cell retention of fructose in cultures incubated with 25
mM and 50 mM fructose by 54% and 62.9%, respectively. The erythritol concentrations in Caco-2 cells incubated with 25 mM and 50 mM fructose were 0.018% and 0.013% of the erythritol present in the medium, respectively.
Experiment 2
In the experiment 2, the inhibitory effect of erythritol on fructose transport was still observed, in a dose-dependent fashion (Figure 5.4). However, the degree of
inhibition of fructose transport by erythritol in Caco-2 cells in the experiment 2 was not
as high as that in experiment 1. Fructose transport to the basolateral compartment
decreased by only 6% in the presence of equimolar amount (25 mM) of erythritol, and
11.1% in the presence of 50 mM erythritol.
Experiment 3
Fructose transport to the basolateral compartment was 1.69 % of the fructose present in
the test medium in the absence of erythritol. The addition of 12.5 mM and 25 mM
erythritol to the test medium containing 25 mM fructose lowered fructose transport to
1.65 % and 1.52 % of the fructose in the medium, respectively. Fructose transport was
significantly reduced by the presence of 25 mM erythritol in Caco-2 cells incubated with
medium lacking glucose in the apical chamber the day before the experiment, but the
degree to which erythritol inhibited glucose secretion was not high (Figure 5.5), 9%
116 reduction in fructose transport in the presence of equimolar amount of fructose and
erythritol. However, erythritol transport to the basolateral compartment at both 12.5 mM
and 25 mM erythritol was 0.3% of the erythritol added to the apical compartment.
Experiment 4
The inhibitory effect of erythritol on fructose secretion which was observed with 25
mM fructose and erythritol was not observed with 1 mM fructose and erythritol (Figure
5.6). Erythritol transport to the basolateral compartment was dose-dependent with 1 mM fructose. The results of a phenol red assay for the experiments ranged from 0.02 to
0.05%/hr/cm2, suggesting the intact tight junctions.
Discussion
Our previous in vivo study demonstrated greatly increased carbohydrate
malabsorption after the intake of fructose and erythritol mixture. However, the source of
carbohydrate was not identified in the study. The present in vitro data using Caco-2 cells
suggested that erythritol inhibited fructose secretion in dose-dependent fashion. In
contrast, erythritol transport from apical to basolateral compartment was not affected by
fructose.
Fructose transport was time-dependent in the pilot study. The fructose flux curve from the apical and the basolateral compartment was unchanged after a 30 min incubation period, and therefore Caco-2 cells were incubated for 30 min for subsequent
experiments.
117 In experiment 1, fructose transport to the basolateral compartment and fructose concentration in Caco-2 cells were inhibited by the addition of 25 mM erythritol when 25
mM and 50 mM fructose were added to the apical compartment. However, erythritol flux and erythritol quantity in cells were not affected by fructose, suggesting the potential for paracellular transport of erythritol.
In experiment 2, fructose failed to inhibit erythritol transport, and erythritol flux
was dose-dependent, suggesting that erythritol might be transported paracellularly.
Concentrations of fructose and erythritol in cells were inconclusive due to large variations within treatment. As shown in figure 5.3, an inhibitory effect of erythritol on
fructose transport was observed.
The goal of experiment 3 was to simulate our human study, in which subjects
fasted overnight the day before measurements of carbohydrate absorption following
treatment. Hence, Caco-2 cells were incubated with glucose-free DMEM the day before
the experiment. The consistent inhibitory effect of erythritol on fructose transport was
observed in the experiment 3.
Interestingly, we found that fructose transport was not inhibited by erythritol when fructose concentration was low (1 mM), whereas erythritol inhibited fructose transport when apical concentration was high (25 mM). Fructose is known to be transported by facilitated diffusion, employing two known transporters. On the apical side, GLUT5 is involved in fructose transport (11). In contrast, GLUT2 is located at the basolateral side (18). GLUT5 has high specificity for fructose. Recently, Kellett et al. proposed the apical GLUT2 hypothesis (19). Fructose uptake in the small intestine of wild-type and GLUT2-null mice fed low carbohydrate (2.5 % dextrose) diets was very
118 similar, indicating GLUT5-mediated transport at low concentration of fructose. On the
contrary, fructose uptake was 5.7-fold higher in wild type mice fed high fructose diets
(65% fructose) compared with that with the low carbohydrate diet (P < 0.001). In
addition, fructose absorption rate was different between wild- type and GLUT2-null mice
fed high fructose diets. Fructose transport was reduced by 60% in GLUT2-null mice
compared with wild-type mice, suggesting that reduced fructose transport in GLUT2-null
mice was caused by lack of GLUT2 in the jejunum of the mice. Hence, GLUT5 is
saturated with a low luminal fructose concentration, whereas GLUT2 in the brush border
is active with high luminal concentrations of fructose. Our data were consistent with the
findings of Kellett et al. (19) suggesting that erythritol and fructose may share a common
transporter (possibly GLUT2) in the intestinal cells. However, it could not be determined
from this study where the specific interaction between fructose and erythritol takes place.
Although we observed that erythritol inhibited fructose absorption both in vivo
and in vitro, the impairment of fructose secretion by erythritol that we observed in vitro
was modest compared with the degree of fructose malabsorption that we observed in vivo.
Whereas erythritol reduced fructose absorption by 6-24% in Caco-2 cells, breath
hydrogen production (an indicator of carbohydrate malabsorption) by subjects receiving
fructose in the presence of erythritol was doubled compared with subjects receiving
fructose only. The possible explanations for why a greater impairment of fructose
secretion was not observed in Caco-2 cells are the use of different concentrations of fructose and erythritol that we used in vivo, and an in vivo factor, such as gastric emptying rates. In vivo, healthy subjects received a mixture of each 556 mM of fructose and erythritol (50 g fructose and 33.3 g erythritol in 500 mL water), of which
119 concentrations are toxic to Caco-2 cells. It is difficult to measure luminal concentrations
of fructose and erythritol after intake of a meal/drink containing fructose and erythritol in
humans. One study reported that fructose concentration in the lumen was 113 mM after
intake of 150 mM fructose concentration (approximately 75% of fructose present in the solution reached the lumen) (20). Hence, we used 25 mM and 50 mM of fructose and erythritol, causing modest inhibition of erythritol on fructose absorption. The fructose concentrations used in this study (25 mM and 50 mM) are the same concentrations used in previous studies that measured intestinal GLUT5 expression by high concentrations of fructose in Caco-2 cells (14, 15). Gastric emptying rate is known to affect the rate of digestion and absorption of carbohydrates in humans. However, there was no gastric emptying phase in vitro.
The fact that erythritol transport was not affected by fructose suggests that erythritol may be transported paracellularly. However, the mechanism by which erythritol inhibited fructose was not identified in this study because the experimental design provided descriptive, not mechanistic, results. Future studies are needed to identify if the
interaction between erythritol and fructose occurs transcellularly (possibly involving a
transporter) and/or paracellularly. To determine if GLUT2 is associated with absorption
between erythritol and fructose, the secretion of fructose and erythritol into the
basolateral compartment could be measured after high concentrations of fructose and
erythritol are added to the test medium in the presence and absence of phloretin, a
GLUT2 inhibitor. Also, measuring GLUT2 expression at the apical and basolateral
compartments will indicate if the apical GLUT2 hypothesis can be applied to fructose
and erythritol transport. In addition, the effect of erythritol on the intestinal transport of
120 other carbohydrates, such as glucose, should be investigated. If erythritol and fructose
share GLUT2 in the small intestine, erythritol could affect the absorption of glucose because a high concentration of glucose triggers GLUT2 translocation from the basolateral to the apical compartment. If erythritol inhibits glucose absorption the way it blocks the absorption of fructose, the presence of erythritol with glucose-containing foods may cause lowering of blood glucose level, which will benefit people with diabetes.
In summary, in Caco-2 cells, fructose transport was inhibited in the presence of erythritol at high doses (25 mM) of fructose and erythritol in a dose-dependent manner.
However, erythritol transport was not affected by fructose. This in vitro result supports
the previous findings of the in vivo study conducted at our lab, the possibility of inhibited fructose absorption by erythritol.
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123
350 b 300 250 b 200 a 150 100 a nm oles/well 50 0 0 102030405060 Time (min)
Figure 5.2. Transport of fructose into the basolateral compartment for different length of incubation. Values are means ± SE, n=3. Values with different superscript letters are significantly different (P < 0.05).
124
1400 c d 1200
1000 a 800 b Fru 600 Ery
nmoles/well 400 200 0 25 mM Fru 25 mM Fru + 50 mM Fru 50 mM Fru + 25 mM Ery 25 mM Ery
Figure 5.3. Transport of fructose and erythritol into the basolateral compartment at high doses of fructose and erythritol. Fru=fructose, Ery=erythritol. Values are means ± SE, n=3. Values with different superscript letters are significantly different (P < 0.05).
125
900 ac c 800 ad bd 700 bd 600 500 Fru 400 D Ery 300 C nmoles/well 200 B 100 A 0 25 mM Fru 25 mM Fru 25 mM Fru 25 mM Fru 25 mM Fru + 12.5 mM + 25 mM + 50 mM + 75 mM Ery Ery Ery Ery
Figure 5.4. Transport of fructose and erythritol into the basolateral compartment after phenol red assay was conducted the day before the experiment. Fru=fructose,
Ery=erythritol. Values are means ± SE, n=3. Values with different capital letters are significantly different from erythritol secretion into the basolateral compartment (P <
0.05). Values with different small letters are significantly different from fructose
secretion into the basolateral compartment (P < 0.05).
126
1000 900 a a Fru b 800 Er y 700 600 500 400 nmoles/well 300 200 B A 100 0 25 mM Fru 25 mM Fru + 12.5 25 mM Fru + 25 mM Er y mM Er y
Figure 5.5. Transport of fructose and erythritol into the basolateral compartment after apical side was pre-treated with glucose-free DMEM the day before the experiment.
Fru=fructose, Ery=erythritol. Values are means ± SE, n=3. Values with different capital letters are significantly different from erythritol transport into the basolateral compartment (P < 0.05). Values with different small letters are significantly different from fructose transport into the basolateral compartment (P < 0.05).
127 160 D 140
120
100 Fru 80 C Ery
nmoles/well 60
40 B
20 A A 0 1mM 1mM 1mM 1mM 1mM 1mM Ery Fru Fru + Fru + Fru + Fru + 1mM 5mM 10mM 20mM Ery Ery Ery Ery
Figure 5.6. Transport of fructose and erythritol to the basolateral compartment at low doses of fructose and erythritol. Fru=fructose, Ery=erythritol.Values are means ± SE, n=3.
Values with different capital letters are significantly different (P < 0.05).
128 CHAPTER 6
CONCLUSIONS
In this dissertation, the glycemic and insulin index of raisins (half of the
carbohydrate of which is fructose) were measured. In addition, both in vivo and in vitro
studies were investigated to determine the effect of erythritol on fructose absorption.
The glycemic index of raisins was not different between healthy sedentary subjects,
athletes, and people with impaired glucose tolerance, but was somewhat higher in the athletes. The insulin index of raisins was also not significantly different among different
groups. However, our results contradict other findings, in which athletes had lower
glycemic index of breakfast cereals than sedentary people. Thus, further studies are
needed to evaluate the effect of training status on glycemic index of foods. Also, people
with impaired glucose tolerance (50.0 ± 2.6 y) were older than subjects in other groups
(25.7 ± 1.3 y and 23.1 ± 1.0 y for healthy sedentary subjects and athletes, respectively) in
our study. Although subjects’ age and glucose tolerance status in that group did not affect
the glycemic and insulin index of raisins, future studies can be conducted to measure a
glycemic and insulin index of a test food with healthy sedentary subjects and age-
matched people with impaired glucose tolerance.
129 The simultaneous ingestion of an equimolar amount of erythritol (a four carbon
sugar alcohol) and fructose resulted in higher breath hydrogen excretion, lower serum fructose levels, and significantly increased serum erythritol levels compared with fructose
only. It appears that erythritol was absorbed at the expense of fructose, but a limitation of
this study was that there was no erythritol-only beverage in the experiment. Hence, the identity of the malabosorbed carbohydrate, whether fructose or erythritol, in the fructose-
erythritol treatment could not be determined via the breath hydrogen technique that was
employed.
To better identify the source of malabsorbed carbohydrate, an in vitro study using
radiolabeled fructose and erythritol was conducted in a Caco-2 cell model. It was
observed that erythritol inhibited intestinal fructose transport at high doses (25 mM) of
fructose and erythritol. However, the degree to which erythritol inhibited fructose
absorption in vitro was not as great as in vivo, possibly due to the use of lower doses of fructose and erythritol in vitro versus the higher luminal concentrations that were probably present in our earlier in vivo study. The mechanism by which erythritol inhibits intestinal fructose absorption was not determined in the in vitro study. Future research is necessary to identify the mechanism by which erythritol inhibits fructose absorption, and also to investigate the effect of erythritol on intestinal absorption of other carbohydrates, such as glucose and galactose. In addition, is needed be determined.
In conclusion, this study demonstrated that high fructose containing food/solution
has low-moderate glycemic and insulin responses, but gastrointestinal distress was
observed by the inhibited fructose absorption in the presence of erythritol. The use of
sugar alcohols in food products for diabetes and weight loss has been increasing due to
130 their lower energy content and glycemic responses compared with sugar. However, there
are few studies that have examined the interaction of sugar alcohols and other foods components which may affect carbohydrate absorption, gastrointestinal symptoms and GI values. Further research is warranted.
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151
APPENDIX A
INSTUTUTIONAL APPROVAL LETTER
FOR THE RAISIN STUDY
152
153
APPENDIX B
RECRUITMENT FLYER
FOR THE RAISIN STUDY
154 Research Subjects Needed
Research subjects are needed to participate in an experiment designed to compare the blood glucose (sugar) and insulin responses to raisins and glucose solution (sugar beverage) in 3 different groups as follows: 1) healthy non-exercising group, 2) competitive athletes, and 3) persons with a reduced ability to use sugar as an energy source. Potential subjects described as follows:
• Males or non-pregnant, non-lactating females • Group 1: healthy non-exercising adults aged 18-35 years without a reduced ability to use sugar as an energy source • Group 2: competitive athletes aged 18-35 years without a reduced ability to use sugar as an energy source • Group 3: persons with a reduced ability to use sugar as an energy source aged 18-65 years. • No use of tobacco products • No history of gastrointestinal disorders • No use of medications (acetaminophen, salicylates, diuretics, etc) that could interfere with nutrient absorption, metabolism, excretion, or stomach motility.
There will be a total of two 2-3-hour blood glucose tests that subjects will be required to complete, one for raisins and glucose solution (sugar solution). For each test, the subject must fast overnight for at least 10 hours. Upon the subject’s arrival, a finger stick blood sample will be obtained for measurements of glucose and insulin. Subjects will then be given either a 2-ounce (64 g) portion of raisins or a 10-ounce (300ml) portion of glucose solution as well as 8 oz of tap water. The subject will not be allowed to eat anything else for the 2-hour period after each test meal. Blood glucose and insulin (finger sticks) will be measured at the following intervals: baseline (immediately before the test meal), and at 15, 30, 45, 60, 90, and 120 minutes following the start of the test meal consumption. For persons with a reduced ability to use sugar as an energy source, additional blood samples at 150 and 180 minutes will be obtained. Subjects will be required to remain in the laboratory for the duration of the 2-3 hour test period. Participants will be paid $50 for proper completion of the entire study. If you meet the above criteria and would like to participate in the study or have questions about the study, please contact: Yeonsoo Kim 219 Campbell Hall 1787 Neil Avenue Columbus, OH 43210 (614) 292-1239 [email protected] [email protected] 292-1239 (614) Yeonoso Kim, MS [email protected] 292-1239 (614) Yeonoso Kim, MS [email protected] 292-1239 (614) Yeonoso Kim, MS [email protected] 292-1239 (614) Yeonoso Kim, MS [email protected] 292-1239 (614) Yeonoso Kim, MS [email protected] 292-1239 (614) Yeonoso Kim, MS [email protected] 292-1239 (614) Yeonoso Kim, MS [email protected] 292-1239 (614) Yeonoso Kim, MS
155
APPENDIX C
SCREENING FORM
FOR THE RAISIN STUDY
156 Screening Questionnaire
This form contains information that will be used to determine your eligibility to participate in a study entitled “determination of the glycemic and insulinemic responses to raisins” Please PRINT all information in the white boxes. Members of the study staff will fill in shaded areas.
Demographic Data
______FIRST NAME M.I. LAST NAME SCREEN DATE
______SOCIAL SECURITY NUMBER
______STREET ADDRESS HOME PHONE NUMBER
______CITY STATE ZIP WORK PHONE NUMBER
______DATE OF BIRTH AGE EMAIL
RACE: American Indian or Alaska Native Asian Black or African American Native Hawaiian or Other Pacific Islander White Other: ______
ETHNICITY: Hispanic Non-Hispanic
GENDER: Male Female
Anthropometric Data FILLED IN BY STUDY STAFF
______HEIGHT (cm) WEIGHT (kg) BMI (kg/m2)
Fasting Blood Glucose Test
When did you begin fasting last night? ______pm
______TIME OF SAMPLE (HH:MM) # HOURS SINCE LAST MEAL (HH:MM) FASTING BLOOD GLUCOSE (mg/dL)
157 Health Screening Information
1. Did you voluntarily sign an informed consent form prior to NO participation in this study? YES
2. Did you receive a copy of your signed informed consent NO form? YES 3.Are you currently pregnant? (MALES – mark N/A) N/A NO YES Pregnancy test result: NEG (-) POS (+)
______STAFF INITIALS DATE
1. Are you at least 6 weeks post-partum and non-lactating? N/A NO (MALES and FEMALES WHO HAVE NOT HAD A CHILD YES mark N/A)
2. Have you been diagnosed with impaired glucose NO tolerance? YES
If yes, when? ______Month Year
Duration of having impaired glucose tolerance
______Month Year
3. Do you have any active metabolic or gastrointestinal NO diseases? YES If yes, please list: ______
______
4. Do you have any clotting or bleeding disorders? NO If yes, please List:______YES
5. Do you have any known food allergies or intolerances NO (excluding lactose intolerance)? YES If yes, please list:______
______
6. Have you had an infection requiring hospitalization, NO surgery, or corticosteroid treatment in the last 3 months? YES 158
7. Have you fainted or had other adverse reactions in NO response to blood collection in the past? YES
8. Are you willing to fast overnight for 10 hours before NO treatment visits 1 and 2? YES
9. Are you willing to consume the study product within a 10- NO minute period? YES
10. Are you willing to abstain from exercise (defined as any NO moderate exertion that is beyond normal, daily activity YES levels) 24 hours prior to treatment visits 1 and 2, and minimize activity during the test?
11. Are you willing to remain awake for the duration of NO treatments within a 2-3 hour period? YES
12. Are you willing to continue fasting for 2-3 hours after NO consuming product during treatment visits 1 and 2? YES
13. Following the start of a test meal, are you willing to NO undergo blood glucose testing via finger stick YES method 7-9 times within a 2-3 hour period? You must remain in the laboratory during this period. 14. Are you willing to complete the necessary forms and NO questionnaires and keep accurate notes and records for YES all study visits?
159 15. Are you taking any supplements on a regular basis? NO YES (If YES, please list below.) Start Stop Dose Form Frequency Reason date date (mg, units, (liquid, pill, (per day, week, etc) injectable, etc) etc)
160
16. Are you currently on any medications? NO YES (If YES, please list below.) Medication Start Stop Dose Form Frequency Reason (list brand, if known) date date (mg, units, (liquid, pill, (per day, etc) injectable, etc) week, etc)
Physical Activity Information
1. Considering a 7-day period (week), how many times on the average do you do the following kinds of physical activity for more than 15 minutes while at work, as part of house or yard work, to get from place to place, and in your spare time for recreation, exercise, and sport?
TARGET ACTUAL PERCENT A) Strenous Physical Activity (Heart beats 4 rapidly) For example, running, jogging, hockey, football, soccer, squash, basketball, cross country skiing, roller skating or blading, vigorous swimming, vigorous long-distance biking B) Moderate Physical Activity (Not 10 exhausting) For example, fast walking, baseball, tennis, easy bicycling, volleyball, badminton, easy swimming, downhill skiing, popular and folk dancing C) Mild Physical Activity (Minimal effort) 20 For example, yoga, archery, fishing from river bend, bowling, horseshoes, golf, snow- mobiling, easy walking
2. Considering a 7-day period (a week), how often do Oftern you engage in any regular activity long enough to Sometimes work up a sweat (heart beats rapidly)? Rarely/Never
3. For how long have you been active at this level? ______day(s) ______month(s) ______year(s)
4. Do you exercise regularly? (If yes, please complete questions 5-9) NO YES
5. What type of endurance exercise do you typically running perform? cycling swimming rowing other______
161 6. How many years have you been en endurance 1-2 years athlete? 3-4 5-6 7-8 9+
7. How many competitions have you participated in 1 during the past 6 months? 2 3 4 5+
8. During the competitive season, how many hours do 1-3 hours you normally train per week? 4-6 7-9 10-12 13+
9. How many hours are you currently training per week? 1-3 hours 4-6 7-9 10-12 13+
I agree that the information I provided for this screening visit is true and accurate to the best of my knowledge.
______SUBJECT’S SIGNATURE DATE SIGNED
______STAFF’S SIGNATURE DATE SIGNED
162
APPENDIX D
TREATMENT VISIT DATA COLLECTION FORM
FOR THE RAISIN STUDY
163 Treatment Visit Data Collection Determination of the Glycemic and Insulinemic Responses to Raisins
Good Morning! Thank you for participating in our study. This form will provide you with directions and serve as a data collection sheet in completing your visit. Please hand in your food record, write down your arrival time in the space provided, and let a member of the study staff measure your weight. Then, rest while you fill out the following Check-In information. A staff member will take your blood pressure, pulse, and temperature at the end of the 30-minute rest period.
C H E C K - I N 30-minute rest period ends Date: Arrival time __ __:__ __ am __ __:__ __ am ____/____/2004 1. Have you experienced any unusual symptoms or medical problems since your last visit? If yes, please describe: NO ______YES Did these symptoms or problems cause you to seek medical attention? If yes, specify name(s) of medical professional(s): ______NO ______Please describe any action taken (started meds, admitted to hospital, YES etc.): ______
2. Have you made any changes to your daily medications or supplements since NO your last visit? If yes, please describe: YES ______
3. Have you exercised during the last 24 hours? NO
YES 4. Did you fast at least 10 hours last night? If no, please explain: NO ______YES
am 5. When did you begin Date: Time: __ __:__ __ pm fasting? ____/____/2004 F I L L E D I N B Y S T U D Y S T A F F Blood Pressure: Pulse: Temperature: Weight:
______.__ kg ______/______bpm ______. __°F
Staff Initials ______
164
D A T A C O L L E C T I O N I N S T R U C T I O N S After the 30-minute rest period, you are ready to begin collecting data. You have a Station with a timer, access to a wall clock, tubes for collecting blood.
Blood Draw: On a rack at your station, set up your blood collection tubes in order of the time points on their labels (000, 015, 030, 045, 060, 090, and 120min). A nurse or emergency medical technician will be drawing your blood at specified intervals.
Corrections If you make an error on this form, please put one horizontal line through the error, write the correct information above or beside the error, and initial and date the change. Please record everything using an ink pen.
It is important that your blood be tested at precise intervals, so when your timer goes off, immediately reset your clock. Make sure the study staff collects your blood sample and records the time it was collected within the +5 min window. All times should be reported using the wall clocks in the lab and metabolic kitchen area. Each time point that you collect a blood is in reference to the time at which you began drinking the study beverage (i.e. if you begin drinking the study beverage at 7:45 a.m., your 15 minute time would be 8:00 a.m.)
After baseline samples are taken in the lab, come back to the metabolic kitchen, start your timer right before you consume your study product, and then consume your study beverage.
The following is an outline of steps that you will follow in your collection of data. Complete the steps in the order they are written. Shaded areas will be filled in by the study staff.
0 0 0 M I N U T E S ( B A S E L I N E )
am pm Finger stick __ __:__ __
Walk to metabolic kitchen.
Set timer for 15 MINUTES.
Time you started drinking study product. __ __:__ __ am Consume the study product within 10 minutes. pm
Time you finished study product. __ __:__ __ am pm
Wait for timer to go off. 0 1 5 M I N U T E S
Set timer for 15 MINUTES.
Finger stick __ __:__ __ am pm
165
Wait for timer to go off. 0 3 0 M I N U T E S
Set timer for 15 MINUTES.
Finger stick __ __:__ __ am pm
Wait for timer to go off. 0 4 5 M I N U T E S
Set timer for 15 MINUTES.
Finger stick __ __:__ __ am pm
Wait for timer to go off.
0 6 0 M I N U T E S ( 1 H O U R )
Set timer for 30 MINUTES.
Finger stick __ __:__ __ am pm
Wait for timer to go off. 0 9 0 M I N U T E S
Set timer for 30 MINUTES.
Finger stick __ __:__ __ am pm
Wait for timer to go off. 1 2 0 M I N U T E S ( 2 H O U R S )
Finger stick __ __:__ __ am pm
166 E N D O F S T U D Y V I S I T
Thank you for completing this phase of the study!!!
Please sign:
Subject: ______Date:______
Study staff: ______Date:-
______
167
APPENDIX E
WESTERN INSTITUTIONAL REVIEW BOARD
APPROVAL LETTER FOR THE ERYTHRITOL AND FRUCTOSE STUDY
168
169
170
APPENDIX F
RECRUITMENT FLYER FOR THE ERYTHRITOL AND FRUCTOSE STUDY
171 Volunteers Needed
Volunteers are being recruited to participate in a research study designed to evaluate the effect of erythritol, a natural sweetener, on fructose absorption in healthy adults, via non- invasive breath hydrogen test. Blood sugar, insulin responses, and gastrointestinal tolerance will also be evaluated.
Potential subjects described as follows: Age 18-75 years Body weight within approximate normal range Males or non-pregnant, non-lactating females No history of diabetes No antibiotic use within the last three weeks No use of tobacco products Hydrogen producer (a screening test to assess the ability to produce hydrogen gas)
Volunteers will be screened for the ability to produce hydrogen gas using a non-invasive breath hydrogen test. The volunteers will be given about 2 tsp of lactulose, a non-absorbable sugar, in 8 fluid ounces of water. Breath samples will be collected at baseline (before drinking a beverage containing lactulose) and each hour up to 8 hours after drinking a beverage, and a 1-hour visit to the lab (219 Campbell Hall) is also required. Upon the entry into the study, participants will complete 3 study treatments over 3 to 6 weeks. Each treatment will require an overnight fast, consumption of a study beverage containing fructose, glucose, and/or erythritol, followed by collection of blood and breath samples. Study visits run from 7:00 a.m. until approximately 11:00 a.m. in the laboratory (219 Campbell Hall) with additional collection of breath samples at home until approximately 4:00 p.m. More information will be provided at screening. Please feel free to contact the study coordinator if you have any questions. Subjects will be paid up to $320 for completion of the study and including $20 for the breath hydrogen screening test.
Principal Investigator: Study coordinator: Steve Hertzler, PhD, RD Yeonsoo Kim, MS 341 Campbell Hall 219 Campbell Hall 1787 Neil Ave. 1787 Neil Ave. Columbus, Ohio 43210 Columbus, Ohio 43210 (614) 292-5575 (614) 292-1239 [email protected] [email protected]
SRDB-14, Version 1, 11/20/03
[email protected] (614) 292-1239 Kim, MS Yeonsoo [email protected] (614) 292-1239 Kim, MS Yeonsoo [email protected] ( Kim, MS Yeonsoo [email protected] (614) 292-1239 Kim, MS Yeonsoo [email protected] (614) 292-1239 Kim, MS Yeonsoo [email protected] (614) 292-1239 Kim, MS Yeonsoo [email protected] (614) 292-1239 Kim, MS Yeonsoo [email protected] (614) 292-1239 Kim, MS Yeonsoo 614) 292-1239
172
APPENDIX G
SCREENING QUESTIONNAIRE FOR THE ERYTHRITOL AND FRUCTOSE
STUDY
173 Screening Questionnaire
This form contains information that will be used to determine your eligibility to participate in the protocol “an evaluation of the effect of erythritol on fructose absorption in healthy adults.” Please PRINT all information in the boxes.
______FIRST NAME M.I. LAST NAME SCREEN DATE
______STREET ADDRESS HOME PHONE NUMBER
______CITY STATE ZIP WORK PHONE NUMBER
______DATE OF BIRTH AGE EMAIL
RACE: White Black or African American Asian Native Hawaiian or Other Pacific Islander American Indian or Alaska Native Other: ______
ETHNICITY: Hispanic Non-Hispanic
GENDER: Male Female
Anthropometric Data FILLED IN BY STUDY STAFF
______HEIGHT (cm) WEIGHT (lbs) WEIGHT (kg) BMI (kg/m2)
______EER (kcal) # ENSURE BARS WAIST CIRCUMFERENCE (in, if BMI > 28)
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Health Screening Information
1. Are you currently pregnant? (MALES – mark N/A) N/A NO YES (ALL FEMALES AT CHILD BEARING AGE HAVE TO TAKE PREGNANCY TEST)
Pregnancy test result: NEG (-) POS (+)
______STAFF INITIALS DATE
2. Are you at least 6 weeks post-partum and non- N/A NO YES lactating? (MALES and FEMALES WHO HAVE NOT HAD A CHILD – mark N/A)
3. Are you 18 - 75 years of age? NO YES
4. Do you have diabetes mellitus or glucose NO YES intolerance?
5. Do you have any active metabolic or NO YES gastrointestinal diseases? If yes, please list: ______
6. Do you smoke? NO YES
7. Do you have any known food or drug allergies or NO YES intolerances (excluding lactose)? If yes, please list: ______
8. Have you experienced some adverse reaction in NO YES response to blood collection before?
9. Have you had antibiotics in the last 3 weeks? NO YES
10. Are you willing to consume Ensure Plus® as your NO YES evening meal on the day prior to testing and controlled lunch on study treatment visit?
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11. Are you willing to abstain from exercise 24 hours prior to NO YES testing and throughout the study visits until the last breath hydrogen sample is collected?
12. Are you willing to consume study beverages containing NO YES fructose, glucose and/or erythritol?
13. Are you willing to remain awake for the duration of the NO YES study visits until the last breath hydrogen sample is collected?
14. Did you voluntarily sign an informed consent form prior to NO YES participation in this study? 15. Following the start of a study beverage, are you willing to NO YES have an indwelling catheter for 3 hours for 9 venous blood draws? Are you also willing to place your hand into a heated hand box to warm your hand before each blood draws? (You must remain in the laboratory during this period.)
16. Are you willing to continue fasting for additional 4 hours NO YES after consuming provided Ensure Plus® lunch with the exception of water until you complete collecting the 8th hour breath sample?
17. Are you willing to complete the necessary forms and NO YES questionnaires and keep accurate notes and records?
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18. Are you taking any supplements on a regular basis? NO YES (If YES, please list below.) Supplement Start Stop Dose Frequency Reason Form (list brand, if known) date date (mg, units, (per day, week, (diagnosis, (liquid, pill, etc) etc) prescription, etc) injectable, etc)
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19. Are you currently on any medications? NO YES (If YES, please list below.) Medication Start Stop Dose Frequency Reason Form (list brand, if known) date date (mg, units, (per day, week, (diagnosis, (liquid, pill, etc) etc) prescription, etc) injectable, etc)
Pre-Visit Ensure Meal Product Selection
You will need to consume the same flavor of Ensure® beverage and the same flavor of Ensure® bar throughout the study. Please sample the products and indicate (circle) your preference for each below.
Beverage Bar
Chewy Maple Cookies ‘n Cinnamo Chocolate Vanilla Strawberry Chocolate Chocolate Cream n Oat Peanut ‘n Raisin
Were you instructed on storage and use of Ensure® NO YES meal products?
I agree that the information I provided for this screening visit is true and accurate to the best of my knowledge.
______SUBJECT’S SIGNATURE DATE SIGNED
______STAFF’S SIGNATURE DATE SIGNED
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APPENDIX H
BREATH HYDROGEN AND FASTING PLASMA GLUCOSE SCREENING FORM FOR THE ERYTHRITOL AND FRUCTOSE STUDY
179 Breath Hydrogen and Fasting Plasma Glucose Screening
This form contains information that will be used to determine your eligibility to participate in the protocol “an evaluation of the effect of erythritol on fructose absorption in nondiabetic healthy adults.” Please PRINT all information in the white boxes. Shaded areas will be filled in by members of the study staff.
______FIRST NAME M.I. LAST NAME SCREEN DATE
Fasting Glucose Sample When did you begin fasting? am pm DATE FASTING BEGAN (MM/DD/YYYY) TIME FASTING BEGAN (HH:MM)
Did you consume only water and the Ensure Plus® beverage and NO YES Bars after 4:00 p.m. as your evening meal last night? If no, please explain: ______
Completed by Study Staff ______TIME OF SAMPLE (HH:MM) HOURS SINCE LAST MEAL (HH:MM) FASTING BLOOD GLUCOSE (mg/dL)
Breath Samples am Time of fasting breath sample __ __:__ __ pm am Time you began drinking lactulose solution __ __:__ __ pm am am Hour 1 breath __ __:__ __ pm Hour 5 breath sample __ __:__ __ pm sample am am Hour 2 breath __ __:__ __ pm Hour 6 breath sample __ __:__ __ pm sample am am Hour 3 breath __ __:__ __ pm Hour 7 breath sample __ __:__ __ pm sample am am Hour 4 breath __ __:__ __ pm Hour 8 breath sample __ __:__ __ pm sample
Completed by Study Staff
Hydrogen Producer NO YES
Produce more Hydrogen than Methane NO YES
I agree that the information I provided for this screening visit is true and accurate to the best of my knowledge. ______SUBJECT’S SIGNATURE DATE SIGNED ______STAFF’S SIGNATURE DATE SIGNED 180
APPENDIX I
TEST VISIT DATA COLLECTION FORM
FOR THE ERYTHRITOL AND FRUCTOSE STUDY
181 Test Visit Data Collection An Evaluation of the Effect of Erythritol on Fructose Absorption in Nondiabetic Healthy Adults
Good Morning! Thank you for participating in our study. This form will provide you with directions and serve as a data collection sheet in completing your visit. Please hand in your food record and any forms from your previous visit, write down your arrival time in the space provided, and let a member of the study staff measure your weight. Then, put your hand in a heated hand box and rest while you fill out the following Check-In information. A staff member will take your blood pressure, pulse, and temperature at the end of the 30-minute rest period.
C H E C K - I N Date: Arrival time 30-minute rest period ends ____/____/2004 __ __:__ __ am __ __:__ __ am
5. Have you experienced any unusual symptoms or medical problems since your last visit? If yes, please describe: ______NO ______Did these symptoms or problems cause you to seek medical YES attention? NO If yes, specify name(s) of medical professional(s): YES ______
______Please describe any action taken (started meds, admitted to hospital, etc.): ______
______
6. Have you made any changes to your daily medications or supplements NO since your last visit? If yes, please describe: ______YES
7. Have you exercised during the last 24 hours? NO
YES 8. Did you consume only water and the Ensure Plus beverage after 4:00 p.m. as your evening meal last night? If no, please explain: ______NO ______YES
am 6. When did you begin Date: ____/____/2004 Time: __ __:__ __ pm fasting? F I L L E D I N B Y S T U D Y S T A F F Blood Pressure: Pulse: Temperature: Weight:
______.__ kg ______/______bpm ______. __°F
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D A T A C O L L E C T I O N I N S T R U C T I O N S
After the 30-minute rest period, you are ready to begin collecting data. You have a station with a timer, access to a wall clock, tubes for collecting blood, a breath sample collection kit, a heated hand box, and a flatus counter.
Blood Draw: On a rack at your station, set up your blood collection tubes in order of the time points on their labels (000, 015, 030, 045, 060, 090, 120, 150, and 180 min). Your hand should keep being placed in a heated hand box during the 3-hour blood collection period. A nurse will be drawing your blood at specified intervals.
Breath Hydrogen Test: Your breath hydrogen tubes are in a bag and are labeled for 0 – 8 hours. Please be sure that each sample is collected in the appropriate tube. You will stay in the lab until you collect your 3rd hour breath sample, and additional breath sample for hours 4-8 will be collected on your own. Please make sure you collect breath sample within the permitted window (±10 min) of each time point, otherwise, you will be rescheduled to repeat the treatment visit. Your expected breath hydrogen collection time will be based on the time to start drink a study beverage.
Flatus Counter You will count total number of rectal gas passage for 8 hours after drinking a study product, using a flatus counter.
Controlled Lunch Meal You will consume Ensure Plus® provided as a controlled lunch after the completion of blood sampling, which approximates one third of your energy needs.
If you make an error on this form, please put one horizontal line through the error, write the Correct information above or beside the error, initial and date the change. A nurse should also initial and date your errors. It is important that your blood be tested at precise intervals, so please immediately record the time and reset your clock when the timer goes off. Please record everything using an ink pen. All times should be reported using the clocks in the lab and metabolic kitchen area.
After baseline samples are taken in the lab, come back to the metabolic kitchen, start your timer right before you consume your study product, and then consume your study beverage.
The following is an outline of steps that you will follow in your collection of data. Complete the steps in the order they are written.
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0 0 0 M I N U T E S ( B A S E L I N E ) am Catheter inserted. Venous blood draw. __ __:__ __ pm
Collect breath hydrogen sample. __ __:__ __ am pm
Walk to metabolic kitchen.
Set timer for 15 MINUTES.
Record time you started drinking study am beverage. __ __:__ __ pm Drink the beverage within 10 minutes.
Record time you finished study beverage. __ __:__ __ am pm
Place your hand in a heated hand box.
Wait for timer to go off. 0 1 5 M I N U T E S
Set timer for 15 MINUTES.
Venous blood draw. __ __:__ __ am pm
Wait for timer to go off. 0 3 0 M I N U T E S
Set timer for 15 MINUTES.
Venous blood draw __ __:__ __ am pm
Wait for timer to go off. 045 MINUTES
Set timer for 15 MINUTES.
Venous blood draw __ __:__ __ am pm
Wait for timer to go off.
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0 6 0 M I N U T E S
Set timer for 30 MINUTES.
Venous blood draw __ __:__ __ am pm
Collect breath hydrogen sample. __ __:__ __ am pm
Wait for timer to go off. 0 9 0 M I N U T E S
Set timer for 30 MINUTES.
Venous blood draw __ __:__ __ am pm
Wait for timer to go off. 1 2 0 M I N U T E S ( 2 H O U R S )
Set timer for 30 MINUTES.
Venous blood draw __ __:__ __ am pm
Collect breath hydrogen sample. __ __:__ __ am pm
Wait for timer to go off. 1 5 0 M I N U T E S
Set timer for 30 MINUTES.
Venous blood draw __ __:__ __ am pm
Wait for timer to go off. 1 8 0 M I N U T E S ( 3 H O U R S )
Set timer for 60 MINUTES.
Venous blood draw. Catheter removed. __ __:__ __ am pm
Take your hand out from a heated hand box.
am Collect breath hydrogen sample. __ __:__ __ pm
Wait for timer to go off.
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E N D O F S T U D Y V I S I T
The laboratory portion of this test visit is complete. You must continue to fast until your last breath sample is collected, with the exception of water and your optional Ensure® Meal. This meal consists of one or two cans of Ensure Plus®. Once you choose your optional meal, it must remain the same for both visits. You must also refrain from exercise and sleep until all breath samples have been collected. You can return your breath samples at your next visit. Please indicate your meal selection below:
Optional 0 cans Flavor (same as pre-treatment meal) Ensure 1 can Meal 2 cans Chocolate Vanilla Strawberry (same for both treatments)
Please remember to take your breath collection kit and the last page of this packet home with you and return it at your next visit.
Thank you for completing this phase of the study!!!
Please sign:
Subject: ______Date:______
Nurse: ______Date:______
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APPENDIX J
FLATUS/BOWEL MOVEMENTS FORM
FOR THE ERYTHRITOL AND FRUCTOSE STUDY
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APPENDIX K
GASTROINTESTINAL TOLERAMCE FACTORS FORM
FOR THE ERYTHRITOL AND FRUCTOSE STUDY
189
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APPENDIX L
EXIT VISIT QUESTIONNAIRE
191 End of Study / Exit Visit Questionnaire
1. Are you stopping your study participation before completing the NO study? YES If yes, please explain why: ______
1. Have you experienced any unusual symptoms or medical NO problems YES since your last visit? If yes, please describe: ______If yes, did the aforementioned symptoms or problems cause you to seek medical attention? NO YES If yes, specify name(s) of medical professional(s):______Please describe any action taken (started meds, admitted to hospital, etc.): ______
2. Are you interested in being contacted for future nutrition NO studies YES by the Department of Medical Dietetics? If yes, how may we reach you? PHONE: ______EMAIL: ______
Please feel free to make any comments about the trial in the space provided below.
______
______
Please sign and date.
______SUBJECT’S SIGNATURE DATE SIGNED
______SCREENER’S SIGNATURE DATE SIGNED
Thanks for participating in this study!
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