Chromium propionate in broilers: effect on insulin sensitivity
M. A. Brooks,∗,1 J. L. Grimes,† K. E. Lloyd,∗ K. Krafka,‡ A. Lamptey,‡ and J. W. Spears∗,2
∗
Department of Animal Science, North Carolina State University, Raleigh, NC 27695-7621; † Prestage
Department of Poultry Science, North Carolina State University, Raleigh, NC 27695-7608; and ‡ Kemin Agrifoods
North America, Inc., Des Moines, IA 50301
ABSTRACT The objective of this study was to evaluate the effects of dietary chromium (Cr), as chromium
propionate, on measures of insulin sensitivity. Liver and
muscle glycogen, and plasma glucose and non-esterified
fatty acid (NEFA) concentrations were used as indicators of insulin sensitivity. In total, 288 newly hatched
male Ross broilers were divided into 4 dietary treatments consisting of 0 (control diet analyzed 0.43 to
0.45 mg Cr/kg), 0.2, 0.4, or 0.6 mg supplemental Cr/kg
diet, resulting in 4 treatments with 9 replicate pens
per treatment containing eight birds per pen. At d 21,
2 birds per cage were removed based on the greatest
deviation from pen mean BW, resulting in each pen
containing 6 birds for the final analyses. Final BW
were taken on d 40, and on d 42 two birds from each
pen were sampled for plasma NEFA, glucose, and muscle and liver glycogen determination at the initiation
and termination of a 22 h fast. The remaining 2 fasted
birds were sampled after a 30 min refeeding period. No
differences were observed in feed intake, BW gain, or
feed efficiency on d 21 or d 40. Liver glycogen tended
(P = 0.10) to be greater in Cr-supplemented chicks
in the fed state, and muscle glycogen concentrations
tended (P = 0.07) to be greater in Cr-supplemented
chicks compared with controls following fasting and
refeeding. Plasma glucose concentrations were not affected by dietary Cr in the fed, fasted, or refed state.
Plasma NEFA levels were not affected by treatment
in fed or fasted birds. However, plasma NEFA concentrations were lower (P < 0.01) in chicks supplemented
with Cr than in controls following fasting and refeeding, suggesting that Cr increased insulin sensitivity. No
differences were detected among birds supplemented
with 0.2 or 0.4 mg Cr/kg, and among those receiving 0.4 or 0.6 mg Cr/kg. Results of this study indicate that Cr propionate supplementation of a control
diet containing 0.43 to 0.45 mg Cr/kg enhanced insulin
sensitivity.
Key words: chromium, insulin, broiler, glucose
2016 Poultry Science 95:1096–1104
http://dx.doi.org/10.3382/ps/pew018
INTRODUCTION
Glucose metabolism in avian species differs considerably from mammals. Blood glucose concentrations
are much higher in birds while insulin levels are lower
(Braun and Sweazea, 2008). Birds are also considered
to be less sensitive to insulin than mammals (Scanes,
2009). Chromium (Cr) is known to enhance insulin
sensitivity in mammals (Vincent, 2001), and Cr supplementation has reduced plasma glucose and nonesterified fatty acid (NEFA) concentrations in broilers
(Lien et al., 1999). Heat stress and other types of stress
increase circulating concentrations of corticosterone in
broilers. It is well documented that corticosterone reduces insulin sensitivity in broilers (Zhao et al., 2009).
Studies (Sands and Smith, 1999; Sahin et al., 2002; Zha
et al., 2008) have indicated that Cr supplementation
C 2016 Poultry Science Association Inc.
Received September 11, 2015.
Accepted December 1, 2015.
1
Present address: Oregon Zoo, Portland, OR 97077.
2
Corresponding author: Jerry [email protected]
to practical diets can increase performance of broilers
reared under heat stress conditions.
Inorganic Cr supplementation, at a relatively high
level (20 mg/kg), increased body weight gain, liver
glycogen, and fatty acid synthesis in turkey poults
(Rosebrough and Steele, 1981; Steele and Rosebrough,
1979, 1981). Supplementation with inorganic Cr chloride (20 mg/kg) also increased incorporation of carbon
labeled glucose into liver fatty acids in chicks (Cupo
and Donaldson, 1987). Promoting glycogen and fatty
acid synthesis are well documented effects of insulin.
The present study was conducted to determine the effects of dietary Cr (as Cr propionate) on measures of
insulin sensitivity in broilers.
MATERIALS AND METHODS
Experimental Design
Care, handling, and sampling of birds were approved
by the North Carolina State University Animal Care
and Use Committee. Two hundred and eighty-eight
1096
1097
CHROMIUM AND INSULIN SENSITIVITY IN BROILERS
Table 1. Ingredient composition of starter and grower control
diets.
—— Inclusion rate, % —–
Ingredient
Ground corn
Soybean meal (dehulled)
Fat
Corn-chromium premix
Calcium carbonate
Phosphoric acid
White salt (NaCl)
Sodium bicarbonate
L-lysine
DL-methionine
Threonine
Choline chloride
Mineral premix1
Vitamin premix2
Calculated nutrient analysis
ME, kcal/kg
CP, %
Digestible Lys, %
Digestible Met + Cys, %
Digestible Thr, %
Digestible Try, %
Ca, %
Available P, %
Starter
52.975
36.0
3.0
2.00
2.4
1.8
0.275
0.20
0.375
0.375
0.10
0.20
0.20
0.10
3019
22.4
1.39
0.98
0.86
0.28
1.10
0.56
Grower
57.825
30.0
5.0
2.00
2.2
1.6
0.25
0.25
0.125
0.250
–
0.20
0.20
0.10
3243
19.9
1.03
0.80
0.67
0.25
1.00
0.50
1
Provides per kg of diet: 40 mg Zn (from ZnSO4 ), 60 mg Mn (from
MnSO4 r H2 O), 60 mg Fe (from FeSO4 r H2 O), 10 mg Cu (from
Cu2 (OH)3 Cl Tribasic), 0.30 mg Se (from Na2 SeO3 ), 1.0 mg I (from
Ca(IO3 )2 ), 551 mg Ca (CaCO3 carrier).
2
Provides per kg of diet: vitamin A, 13,200 IU; vitamin D3 , 4,000
IU; vitamin E, 66 IU; vitamin B12 , 0.04 mg; riboflavin, 13.2 mg; niacin,
110 mg; d-pantothenate, 22 mg; vitamin K (from menadione), 4 mg; folic
acid, 2.2 mg; thiamine, 4.0 mg; pyridoxine, 7.9 mg; d-biotin, 0.253 mg;
ethoxyquin, 100 mg; wheat midds, 198.9 mg.
male broilers (Ross 708 Strain; NC State University
Hatchery, Raleigh, NC) were used in this study. Newly
hatched chicks (d 0) were weighed and identified with
numerical neck bands upon delivery to the experimental
facility. Birds were stratified by weight and randomly
assigned to 1 of the 4 treatments to produce similar
average initial BW across treatments. Treatments consisted of 0 (control), 0.2, 0.4, or 0.6 mg supplemental
Cr/kg of diet. Within treatment, birds were stratified
by weight and randomly assigned pen numbers 1 to
9 to produce similar average initial BW across pens
(∼42 g), resulting in 4 treatments with 9 replicate cages
per treatment. During the starter phase (d 0 to 21) 8
chicks were housed in each replicate cage. At the end
of the starter phase, 2 birds per replicate cage were
removed based on deviation from mean BW, and the
6 most uniform chicks in each cage were retained, and
switched to a grower diet for the remainder of the study.
Diets and Housing
Ingredients and also the calculated nutrient composition of the starter and grower diets are shown in
Table 1. The diets were corn-soybean meal-based diets formulated to meet or exceed all nutrient requirements for male broilers (NRC, 1994). Experimental diets were mixed at the North Carolina State University
Feed Mill. One base mix of the starter and grower diets
Table 2. Chemical analysis of control starter and grower diets.
DM, %
CP, % DM
NDF, % DM
Ca, % DM
P, % DM
Mg, % DM
K, % DM
Na, % DM
Fe, mg/kg DM
Zn, mg/kg DM
Cu, mg/kg DM
Mn, mg/kg DM
Starter
Grower
90.2
25.2
12.4
1.23
0.90
0.18
1.17
0.19
236
95
17
87
90.6
23.0
10.4
1.11
0.88
0.17
1.09
0.19
198
78
14
90
was produced. Treatment diets were then prepared by
mixing 2% of a corn-Cr propionate premix with 98%
base mix. Chromium propionate (KemTRACE Cr propionate; Kemin Agrifoods North America, Des Moines,
IA) was mixed with finely ground corn to provide 0, 0.2,
0.4, or 0.6 mg supplemental Cr/kg diet, when added at
2% to the base mix. Analyzed chemical composition of
the control diets is presented in Table 2.
Chicks were housed in heated, thermostatically controlled, PVC coated wire Start Grow Battery Cages
(Alternative Design Mfg., Siloam Springs, AR) with a
23:1 light:dark cycle. The initial temperature set point
was 29.5◦ C when chicks were placed in cages. The room
temperature was then reduced by 2.8◦ C each week until
chicks were 28 d of age. Birds were fed by pen and allowed ad libitum access to feed and water. Body weights
were taken on d 0, 21, and 40, and feed disappearance
was recorded as a measure of feed intake.
Sampling
Weekly samples were taken of the experimental
starter and grower diets. These samples were stored and
composited within diet and feeding period to be used
for later Cr analysis.
On d 42, 2 birds from each pen (fed state) were selected. Pens were sampled as 1 of 9 replicate blocks with
1 pen from each treatment sampled in each block to account for time of termination. Blood was collected via
the wing vein into one tube containing sodium fluoride
for plasma glucose and another tube containing heparin
for NEFA. Tubes were then placed on ice until they
could be transported to the laboratory for processing.
Immediately following blood collection all birds were
euthanized via an IV bolus of 60 mg sodium pentobarbital/kg BW, to prevent loss of muscle glycogen during
harvesting (Edwards et al., 1999). Following termination, samples of breast muscle and liver were extracted
and immediately flash frozen in liquid N2 and stored
at −80 ◦ C for later analysis. The remaining birds were
fasted over a 22 h period, at which point, 2 birds from
each pen (fasted state) were sampled for plasma, muscle, and liver using the replicate block sampling method
described above. Following the 22 h fast, the last 2 birds
in each pen were offered treatment diets for 30 min,
1098
BROOKS ET AL.
after which feed was removed. Refeeding was staggered
by 30 min between each replicate block (4 cages, 1 per
treatment) to allow for blood to be collected between
110 and 130 min following removal of the diets. The
final 2 birds from each pen (refed state) were sampled
for plasma, muscle, and liver as described above.
Laboratory Analysis
Experimental diets were prepared for Cr analysis by
wet ashing with trace metal grade HNO3 (Trace Metal
Grade, Fisher Scientific, Raleigh, NC) using a hot plate
digestion procedure (Lloyd et al., 2010). Chromium
was measured using flameless atomic absorption spectrophotometry (Shimadzu, Model AA-6701F, Kyoto,
Japan) as described by Lloyd et al. (2010). The method
of standard addition was used to remove matrix effects. Each sample plus standard were run in triplicate. Bovine muscle obtained from the National Institute of Standards and Technology (Gaithersburg, MD)
and certified to contain 71 ± 38 ng Cr/g was used as
a reference standard. Composite samples of the control diets were analyzed for chemical components at a
commercial laboratory (Dairy One Forage Laboratory,
Ithaca, NY).
All blood samples were centrifuged at 400 × g for
20 min at 10◦ C and the plasma portion was transferred
to separate tubes and stored at −20◦ C. Plasma glucose
was determined enzymatically using the Glucose (HK)
Assay Kit (Sigma #GAHK-20, Sigma-Aldrich, Inc., St.
Louis, MO) per manufacturer’s instructions. Plasma
NEFA concentration was measured using the WAKO
NEFA-HR(2) Microtiter Assay Kit (HR Series NEFAHR (2), Wako Diagnostics, Richmond, VA) per manufacturer’s instructions. Samples for plasma glucose or
NEFA were reanalyzed when the CV of duplicate samples was greater than 10%.
Glycogen was determined using the glycogen assay
outlined by Passonneau and Lauderdale (1974) as modified by Wende et al. (2007). Briefly, frozen liver or
muscle tissue was fragmented on dry ice and 40 mg of
sample was homogenized in 1 mL of deionized H2 O using a Mini-Beadbeater (Biospec Products, Bartlesville,
OK). The sample was immediately placed in boiling
water for 5 min, after which it was frozen for later
analysis. An aliquot of this extract was reacted with
and without amyloglucosidase (Sigma #A-7420, SigmaAldrich, Inc., St. Louis, MO) in 50 mM Na acetate
(pH 5.5) and 0.2% bovine serum albumin. The sample
was then reacted with glucose reagent (Sigma #G3293,
Sigma-Aldrich, Inc., St. Louis, MO), and the resulting changes in absorption at 340 nm were compared
against a glycogen standard exposed to amyloglucosidase (Sigma #G8876, Sigma-Aldrich, Inc., St. Louis,
MO). Results are presented as glycogen released as glucose, corrected to tissue weight. Samples were reanalyzed for glycogen when: 1) CV of duplicate samples was
greater than 10%, 2) a given glycogen value appeared
Table 3. Analyzed chromium concentrations in experimental
diets.
Supplemental
Treatment
Control
Cr 0.2
Cr 0.4
Cr 0.6
Analyzed Cr, mg/kg
Cr1 mg/kg
Starter
Grower
0.00
0.20
0.40
0.60
0.45
0.66
0.84
1.05
0.43
0.62
0.84
1.07
1
KemTRACE Chromium Propionate (Kemin AgriFoods North America, Des Moines, IA).
to be out of line with other birds in that treatment, or
3) samples needed to be diluted further because they
analyzed above the high standard.
Statistical Analysis
Growth data were analyzed as a completely randomized design and plasma and tissue data were analyzed as
a completely randomized block design using the GLM
procedure in SAS (SAS Institute Inc., 2008). For both
analyses, pen was the experimental unit, and for the
plasma and tissue data, termination block was used
as the blocking variable. Differences among treatments
were determined using single degree of freedom orthogonal contrasts. Comparisons made were: 1) control vs. all
Cr-supplemented treatments, 2) 0.2 vs. 0.4 mg Cr/kg,
and 3) 0.4 vs. 0.6 mg Cr/kg. An outlier test was also
performed on the NEFA data using Proc Reg Residual estimation. The residual or standard error of each
observation was estimated within each treatment by
the difference between the predicted value, based on
the regression equation, and the actual observed value.
Residuals with 5 or more standard errors from zero were
considered outliers and were not used in the statistical
analysis. A pooled analysis was also conducted across
all sampling times to determine the effect of physiological state (fed, fasted, and refed) on each variable. The
model included state, block, treatment, and all possible
interactions.
RESULTS AND DISCUSSION
Chromium Analysis of Diets
Analyzed chromium concentrations in composite
samples of starter and grower diets are shown in
Table 3. The control starter and grower diets analyzed
0.45 and 0.43 mg Cr/kg of diet, respectively. Analyzed
chromium levels for starter and grower diets supplemented with 0.2, 0.4, and 0.6 mg Cr/kg diet were
0.66, 0.62, 0.84, 0.84, 1.05, and 1.07 mg/kg diet, respectively. Analyzed Cr concentrations for diets supplemented with Cr were in line with expected values.
Water samples obtained on the last day of the study
averaged 0.93 ng Cr/mL.
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CHROMIUM AND INSULIN SENSITIVITY IN BROILERS
Table 4. Mortality recorded for male broilers fed diets containing differing levels of supplemental chromium.1
Dietary Cr supplementation2
Diagnosis, bird
0.0 mg/kg 0.2 mg/kg 0.4 mg/kg 0.6 mg/kg
Poor body condition3
Leg abnormalities4
Total mortality
# per treatment
% per treatment5
4
3
4
2
3
3
0
2
7
9.72
6
8.33
6
8.33
2
2.78
1
Initial experimental group contained 288 birds with 72 birds per
treatment.
2
Provided from KemTRACE Chromium Propionate (Kemin AgriFoods North America, Des Moines, IA).
3
Includes dehydration, continued weight loss, and/or loss of body condition.
4
Manifested as either unilateral or bilateral chondrodystrophy.
5
Calculated as total mortality per treatment/total birds per treatment.
Mortality
During the course of the study, several birds were lost
or removed due to either poor body condition or leg
abnormalities. These mortalities have been recorded in
Table 4. Birds were sent for necropsy at the Rollins Animal Disease Diagnostic Laboratory, Raleigh, NC. During necropsy, general body condition was assessed by
visual inspection. Symptoms seen in the birds included
poor feathering, weight loss, and/or leg abnormalities.
In short, 3 birds died of dehydration (one case of which
was secondary to a leg abnormality) in the first week (2
in control, 1 in Cr 0.2), 1 bird died of food aspiration
(Cr 0.2), 10 birds were euthanized due to leg abnormalities (3 in control, 2 in Cr 0.2, 3 in Cr 0.4, and 2
in Cr 0.6), 1 bird suffered from a malformed beak and
was unable to maintain weight (Cr 0.2), and in 6 birds
the cause of death was poor condition with unknown
cause (3 in control, 1 in Cr 0.2, and 2 in Cr 0.4). Thirteen of these deaths (6 in control, 4 in Cr 0.2, and 3
in Cr 0.4) occurred before the 21 d culling; therefore,
these birds accounted for the birds to be removed as per
the experimental design and no loss of duplicate within
replicate occurred. However, the 8 birds lost following
21 d represent a loss of a sampling duplicate, but no
replicate pens were lost during the study. Further, it
was determined that 2 birds (1 in Cr 0.2 and 1 in Cr
0.4) in the study were females. This was confirmed by
dissection during the sampling phase of the study, and
these 2 birds were removed from further analysis.
Performance
Feed disappearance, body weights, gain, and feed efficiency were not affected by treatment (Table 5). Performance results for the starter phase (d 0 to 21) represent
replicate means with 8 birds per cage. For the grower
phase (d 22 to 40) performance results were based on
replicate means of 6 birds per cage. In the present study
birds were housed in batteries in a temperature controlled environment, and probably exposed to minimal
stress. Previous studies also have generally not seen a
growth or feed efficiency response to supplemental Cr in
birds housed in batteries (Kim et al., 1995; Kim et al.,
1996). However, Cr supplementation of broiler diets under heat stress conditions has improved gain and feed efficiency in a number of studies (Sands and Smith, 1999;
Sahin et al., 2002; Zha et al., 2008). Increased concentrations of corticosterone produced during heat or other
types of stress reduces insulin sensitivity (Zhao et al.,
2009), and may explain the performance responses to
Cr in heat stressed birds.
Table 5. Effects of dietary chromium on growth performance of broilers.
Dietary Cr supplementation, mg/kg2
1
Parameter
———————–P-value————————-
0.0
0.2
0.4
0.6
SE
Treatment
Con vs. Cr
0.2 vs. 0.4
0.4 vs. 0.6
Intake, g
0 to 21 d
22 to 40 d
1,129.8
2,862.7
1,138.0
2,858.0
1,122.5
2,842.1
1,151.0
2,846.7
13.45
40.41
0.49
0.98
0.64
0.77
0.42
0.78
0.15
0.94
Body Weight, g
0d
21 d
21 d3
40 d
42.1
862.1
868.1
2,505.4
42.2
850.0
851.2
2,506.2
42.2
856.1
859.2
2,489.2
42.2
838.7
843.0
2,452.5
0.04
13.59
14.56
31.92
0.46
0.66
0.66
0.61
0.35
0.39
0.32
0.54
0.20
0.75
0.70
0.71
0.38
0.37
0.44
0.42
Gain, g
0 to 21 d
22 to 40 d
820.1
1,637.3
807.7
1,654.9
813.9
1,630.0
796.5
1,609.5
13.60
22.76
0.65
0.57
0.38
0.83
0.75
0.45
0.37
0.53
FCR, F:G4
0 to 21 d
22 to 40 d
1.38
1.75
1.41
1.73
1.38
1.75
1.45
1.77
0.027
0.025
0.28
0.73
0.31
0.89
0.46
0.62
0.10
0.53
1
Mean values are based on nine replicate cages of eight chicks per cage during d 0 to 21, and six chicks per cage during
d 22 to 40.
2
Provided from KemTRACE Chromium Propionate (Kemin Agrifoods North America, Des Moines, IA).
3
Mean body weights of chicks after removal of two chicks per cage on d 21.
4
Feed conversion ratio = FCR; F:G = Feed:Gain.
1100
BROOKS ET AL.
Table 6. Time required to sample all birds within each replicate
block.
where supplementation with 0.20 to 0.40 mg Cr/kg diet,
from chromium picolinate, did not affect serum glucose concentrations in non-fasted broilers (Kim et al.,
1995; Lee et al., 2003). However, supplementation
of broiler diets with higher (0.80 mg Cr/kg diet or
greater) chromium concentrations, than those used in
the present study, has reduced serum glucose concentrations in both fed (Kim et al., 1996; Moeini et al.,
2011) and fasted birds (Lien et al., 1999).
Plasma glucose concentrations were lower (P
< 0.001) in fasted compared with fed or refed birds
(Table 8). In previous studies decreased plasma glucose
concentrations were observed in broilers fasted for 12
(Yeh and Leveille, 1970) or 24 h (Edwards et al., 1999).
When birds were refed following fasting, plasma glucose concentrations increased (P < 0.001) above those
measured in the normal fed state. Edwards et al. (1999)
reported that refeeding broilers for 15 min to 6 h, following a 24 h fast, increased plasma glucose concentrations relative to baseline values observed in a fed state.
Plasma glucose was affected by block when samples
were collected following fasting (P = 0.03) and refeeding (P = 0.01) but not when samples were collected in a
fed state (P = 0.61). There are two likely explanations
for the block effects observed. Limited research has indicated diurnal patterns in plasma glucose in both fed and
fasted birds (Davison, 1975). In the present study birds
were bled between 08:16 and 12:34 in the fasted state,
and between 12:59 and 17:19 after feeding. Block effects
could also be explained by some blocks of birds encountering more stress than others. Stress could certainly
affect plasma glucose as well as NEFA concentrations.
The stress level on birds appeared to be minimal in this
study. Birds were removed from cages and bled in the
hallway outside of the bird room. However, stress created by study staff entering the bird room and removing
birds may have varied slightly during the bleeding period. Any differences in plasma glucose due to diurnal
or stress effects would show up as a block effect.
Time (min) required to sample
replicate block2
Replicate block1
Fed3
Fasted3
Refed3
1
2
3
4
5
6
7
8
9
36
33
39
28
29
28
33
21
36
29
31
35
32
30
23
30
25
33
27
24
30
22
29
29
40
45
23
1
Each replicate block consisted of 4 cages with each cage representing
a different dietary treatment.
2
Time measurement started when the first blood sample was taken
from the first bird within a block until the last tissue sample of the last
bird was placed in liquid N2 .
3
Fed birds (0 h feed withdrawal) were sampled on d 42 of the study;
Fasted birds (22 h feed withdrawal) were sampled on d 43 of the study;
Refed birds (22 h feed withdrawal followed by 30 min re-feeding) were
sampled on d 43 2 h after feed removal during the re-feeding period.
Sampling Times
For the fasting period, the time between the initial
feed removal of each block on d 42 until the time of sampling ranged from 21.85 to 22.33 h. The elapsed time
between feed removal and sampling of the refed birds
ranged from 115 to 139 min. The time elapsed exceeded
the initial protocol range of 110 to 130 min for 3 replicate blocks (131, 136, and 139 min). The time required
to sample each replicate termination block is shown in
Table 6. Termination blocks were staggered by 30 min
to account for a 30 min sampling time for each block.
The sampling times ranged from 21 to 45 min. As each
block contained 1 cage from each treatment, any variation within the fed, fasted, or refed state termination
times was equally distributed across treatments and was
accounted for statistically by using a blocking variable.
Plasma Glucose
Plasma NEFA
Plasma glucose concentrations were not affected by
dietary Cr within either the fed, fasted, or refed states
(Table 7). This is in agreement with previous studies
Because NEFA values for some birds appeared to be
outliers, an outlier test was performed on the NEFA
Table 7. Effects of dietary chromium on plasma glucose concentrations in male broilers.1
Dietary Cr supplementation, mg/kg3
0.0
Plasma glucose
Fed
Fasted
Refed
2
202.1
183.4
215.0
0.2
0.4
mg/dL
202.3
205.1
182.7
182.2
217.1
212.5
—————————-P-value————————-
0.6
SE
Treatment
Con vs. Cr
0.2 vs. 0.4
0.4 vs. 0.6
203.5
181.5
216.5
4.11
2.15
2.61
0.95
0.94
0.61
0.74
0.61
0.91
0.64
0.87
0.22
0.79
0.83
0.29
1
Fed birds (0 h feed withdrawal) were sampled on d 42 of the study; Fasted birds (22 h feed withdrawal) were sampled on d 43
of the study; Refed birds (22 h feed withdrawal followed by 30 min refeeding) were sampled on d 43, 2 after feed removal during
the refeeding phase.
2
Mean values are based on two chicks from each of the nine replicate cages per treatment.
3
Provided from KemTRACE Chromium Propionate (Kemin AgriFoods North America, Des Moines, IA).
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CHROMIUM AND INSULIN SENSITIVITY IN BROILERS
Table 8. Effect of physiological state on plasma glucose, plasma NEFA, and liver and muscle glycogen concentrations.
————————–P-value———————
Plasma glucose1
Plasma NEFA2
Liver glycogen3
Muscle glycogen3
Fed
Fasted
Refed
SE
Treatment
Block
State
Trt x State
203.3
207.2
35.30
3.64
182.4
483.4
5.49
3.69
215.3
239.8
26.24
3.88
1.7
8.9
1.10
0.13
0.99
0.70
0.02
0.04
0.53
0.01
0.31
0.65
0.01
0.01
0.01
0.41
0.94
0.14
0.30
0.33
1
Expressed as mg/dL.
Expressed as μ Eq/L.
3
Expressed as mg/g wet tissue.
2
Table 9. Effects of dietary chromium on plasma NEFA concentrations in male broilers.1
Dietary Cr supplementation, mg/kg3
0.0
Plasma NEFA2
Fed
Fasted
Refed
233.6
456.4
271.8
0.2
0.4
μ Eq/L
200.0
186.3
488.1
498.1
229.4
232.4
———————P-value ——————–
0.6
SE
Treatment
Con vs. Cr
0.2 vs. 0.4
0.4 vs. 0.6
208.9
490.8
225.6
18.76
21.51
12.43
0.36
0.54
0.05
0.12
0.16
0.01
0.61
0.75
0.87
0.40
0.81
0.70
1
Fed birds (0 h feed withdrawal) were sampled on d 42 of the study; Fasted birds (22 h feed withdrawal) were sampled on d 43 of the
study; Refed birds (22 h feed withdrawal followed by 30 min refeeding) were sampled on d 43, 2 h after feed removal during the refeeding
phase.
2
Mean values are based on two chicks from each of the nine replicate cages per treatment.
3
Provided from KemTRACE Chromium Propionate (Kemin AgriFoods North America, Des Moines, IA).
date using Proc Reg Residual estimation. Based on
the Proc Reg test 3 birds in the fed state, 2 in the
fasted state, and 1 in the refed state were outliers. Birds
identified as outliers were not included in the statistical analysis of NEFA data. Plasma NEFA concentrations were not affected by treatment in fed or fasted
birds (Table 9). In broilers receiving ad libitum access
to feed, chromium supplementation decreased serum
NEFA concentrations in one study (Kim et al., 1995)
but not in another (Kim et al., 1996). As expected,
plasma NEFA concentrations increased (P < 0.001) by
approximately 2-fold following the 22 h fast (Table 8).
During fasting, lipolysis or release of NEFA from adipose tissue would increase because fatty acids would
become the major energy source used by most tissues.
Two h after refeeding for 30 min, plasma NEFA concentrations were (P = 0.05) affected by overall treatment. Plasma NEFA levels were lower (P = 0.01) in
chicks supplemented with Cr compared with controls
(Table 9). Plasma NEFA concentrations did not differ among broilers supplemented with 0.2 and 0.4 mg
Cr/kg, or among those supplemented with 0.4 and
0.6 mg Cr/kg diet. To our knowledge, this is the first
study in broilers to evaluate the effect of dietary Cr
on circulating NEFA concentrations following fasting
and refeeding. Plasma NEFA concentrations following
refeeding were decreased (P < 0.001) compared with
the fasted state, and tended to be higher (P = 0.06)
than values measured in the fed state (Table 8). Sampling block affected plasma NEFA concentrations in the
fed (P = 0.05) and refed state (P = 0.02) but not when
samples were collected after fasting (P = 0.51). As discussed in the glucose section, the block effects could be
due to diurnal patterns or differences in stress level.
The lower NEFA concentrations following refeeding
in chicks supplemented with 0.2, 0.4, or 0.6 mg Cr/kg
diet are consistent with Cr enhancing insulin sensitivity in liver and adipose tissue. Insulin would be expected to exhibit a larger metabolic role following fasting and refeeding. Fasting for 24 h followed by refeeding for 15 min to 6 h increased plasma glucose and
insulin concentrations in broilers compared with baseline values observed in a normal fed state (Edwards
et al., 1999). In this study, refeeding for 15 min increased plasma glucose by 23% and insulin by 167%
compared with baseline values. This supports the importance of insulin in regulating glucose homeostasis
when birds are suddenly challenged with a glucose load.
Glucagon is the major hormone involved in controlling
energy metabolism of poultry during fasting (Scanes,
2009). During fasting, fatty acid synthesis (lipogenesis)
ceases rapidly, and plasma glucagon concentrations increase, stimulating lipolysis. Refeeding following fasting
rapidly stimulates lipogenesis. Yeh and Leveille (1970)
reported that fasting for 2 h depressed hepatic lipogenesis in broilers by approximately 90%; however, refeeding for as little as 30 min following fasting markedly increased hepatic lipogenesis. Insulin works in opposition
to glucagon and stimulates lipogenesis and other anabolic processes. The major site of fatty acid synthesis
in avian species is the liver (Leclercq, 1984), and glucose
is the major carbon source used for fatty acid synthesis.
Fatty acids synthesized in the liver are primarily used
to make triglycerides that are secreted from the liver as
very low density lipoproteins (VLDL). In a fed state,
the fatty acid portion of triglycerides present in VLDL
would be largely taken up and stored in adipose tissue. This process involves the enzyme lipoprotein lipase
1102
BROOKS ET AL.
hydrolyzing the triglycerides in the endothelial lining
of capillaries in adipocytes to free fatty acids. The
free fatty acids would be taken up by adipocytes and
resynthesized into triglycerides for storage. It has been
demonstrated that synthesis of fatty acids in the liver,
and subsequent transport of the fatty acids to adipose
tissue for storage, occurs rapidly following intraperitoneal injection of 14 C-labeled glucose in chicks (Leveille et al., 1968). Synthesis of triglycerides from fatty
acids in adipose tissue requires glucose for synthesis of
the glycerol 3-phosphate backbone. Insulin is known to
increase glucose uptake by mammalian adipocytes, and
insulin has been shown to enhance glucose uptake as
well as glucose oxidation to CO2 and glucose incorporation into triglycerides in isolated chick adipocytes
(Gomez-Capilla and Langslow, 1977). Adipose tissue in
broilers also has a substantial ability to convert glucose
to glycerol (O’Hea and Leveille, 1969). The amount of
NEFA entering the blood from adipose tissue represents a balance between lipid storage or triglyceride
synthesis and lipolysis. Enhanced glucose uptake by
adipocytes would provide a source of energy and glycerol 3-phosphate for triglyceride synthesis and, thus,
reduce release of NEFA from adipose tissue into the
blood stream.
It is known that poultry are more resistant to insulin than mammals (Scanes, 2009). However, based on
considerable research insulin clearly plays an important
role in glucose homeostasis in poultry. Injecting chicks
either intravenous (Tokushima et al., 2005), intraperitoneal (Akiba et al., 1999), or intracardiac (Vives et al.,
1981) with insulin produces hypoglycemia. Tokushima
et al (2005) injected young chicks (following a 12 h
fast) with porcine insulin or saline simultaneously with
2-deoxy-D-[1-3 H] glucose, a non-metabolized glucoseanalog that is transported by the same carrier as glucose. At 10 min following insulin administration, insulin
increased labeled 2-deoxy-D-glucose uptake by 2-fold in
skeletal muscle and over 4-fold in liver. Insulin administration increased 14 C-labeled glucose incorporation into
fatty acids in liver at 30 min after dosing (Kompiang
and Gibson, 1976). Vives et al. (1981) harvested chicks
(fed state) at different time periods following intracardiac administration of 14 C-glucose, with or without
insulin, and measured incorporation of glucose into
triglycerides. Glucose incorporation into liver triglycerides was observed by 10 min, with maximum 14 Cglucose appearing in liver triglycerides at 60 min following injection. Appearance of 14 C-glucose in triglycerides
was not observed until 30 min post injection in plasma
and 60 min following glucose administration in adipose
tissue. Insulin increased 14 C-glucose incorporation into
triglycerides in liver, plasma, and adipose tissue. Collectively, these studies suggest that insulin stimulates
glucose uptake by muscle, liver, and adipose tissue in
chicks.
Recent studies (Simon et al., 2000; Dupont et al.,
2008) involving administration of anti-insulin serum
also supports a critical role for insulin in chickens. Intra-
venous injection of anti-insulin serum resulted in severe
hyperglycemia in chicks within 30 min (Simon et al.,
2000). In addition to elevated plasma glucose concentration, administration of anti-insulin serum also increased
plasma NEFA concentrations and altered insulin signaling pathways in liver and muscle (Dupont et al., 2008).
Chromium functions in insulin-sensitive tissues by
enhancing the action of insulin. Evidence suggests that
Cr potentiates insulin action by binding to apochromodulin to form chromodulin, a low molecular weight
oligopeptide which activates tyrosine kinase in insulin
receptors (Vincent, 2001). Insulin receptors are less numerous in chicks than in mammals, which may partially explain the higher plasma glucose concentrations
in chicks (Simon, 1988). Recently, supplementation of
broilers with 0.2 mg Cr/kg of diet increased gene expression of insulin receptor and insulin receptor substrate 1 in liver (Liu et al., 2010). These are two of the
critical components of the insulin signaling pathway.
Supplementation of chicks with inorganic CrCl3 at a
relatively high level (20 mg Cr/kg diet) increased incorporation of 14 C-labeled glucose into liver fatty acids
at 30 min following intraperitoneal administration of
labeled glucose (Cupo and Donaldson, 1987).
Glycogen
In the fed state, liver glycogen concentrations tended
to be greater (P = 0.10) in Cr-supplemented chicks
compared with controls (Table 10). Liver glycogen did
not differ among chicks supplemented with 0.2 and
0.4 mg Cr/kg, or among those fed 0.4 and 0.6 mg Cr/kg
diet. Chromium supplementation at a much higher level
(20.0 mg Cr/kg diet) from inorganic CrCl3 , increased
glycogen concentrations and glycogen synthase activity
in liver of fed turkey poults (Rosebrough and Steele,
1981). Lambs supplemented with 0.4 or 0.8 mg Cr/kg
diet, from a high Cr yeast, for 60 d also had higher liver
glycogen concentrations than control lambs (Yan et al.,
2010). In rats, Cr supplementation (1 mg Cr/kg diet)
to a low Cr diet increased liver glycogen synthase activity but did not affect liver glycogen concentrations
(Campbell et al., 1989).
Fasting for 22 h reduced (P < 0.001) liver glycogen
concentrations by approximately 84% compared with
values observed in fed birds (Table 8). The depletion of
glycogen reserves with fasting is consistent with previous studies (Lepkovsky et al., 1960; Leveille, 1966, 1969;
Edwards et al., 1999). Fasting liver glycogen concentrations were not affected by Cr (Table 10). Chromium
supplementation also did not affect liver glycogen concentrations in turkey poults fasted for 48 h (Rosenbrough and Steele, 1981).
Liver glycogen concentrations increased (P < 0.001)
greatly when chicks were refed for 30 min following the
22 h fast (Table 8). However, liver glycogen concentrations in refed birds were still lower (P < 0.001) than
those in fed birds. Edwards et al. (1999) found that
refeeding broilers for 12 h following a 24 h fast increased
1103
CHROMIUM AND INSULIN SENSITIVITY IN BROILERS
Table 10. Effects of dietary chromium on liver and breast muscle glycogen concentrations in male broilers.1
Dietary Cr supplementation, mg/kg3
2
Glycogen
Liver
0.0
0.2
0.4
0.6
————————P-value———————–
SE
Treatment
Con vs. Cr
0.2 vs. 0.4
0.4 vs. 0.6
Fed
Fasted
Refed
————–
31.44
7.04
24.65
mg/g wet tissue ——————
41.55
36.74
31.46
6.26
5.04
3.60
28.46
27.29
24.55
2.61
1.86
2.01
0.03
0.59
0.44
0.10
0.34
0.37
0.20
0.65
0.69
0.17
0.59
0.35
Muscle
Fed
Fasted
Refed
————–
3.38
3.64
3.42
mg/g wet tissue —————3.32
4.06
3.80
3.96
3.84
3.32
3.81
4.36
3.92
0.27
0.20
0.28
0.21
0.16
0.15
0.29
0.78
0.07
0.07
0.68
0.17
0.51
0.09
0.27
1
Fed birds (0 h feed withdrawal) were sampled on d 42 of the study; Fasted birds (22 h feed withdrawal) were sampled on d 43 of the
study; Refed birds (22 h feed withdrawal followed by 30 min refeeding) were sampled on d 43 of the study, 2 h after feed removal during
the refeeding phase.
2
Mean values are based on two chicks from each of the nine replicate cages per treatment.
3
Provided from KemTRACE Chromium Propionate (Kemin AgriFoods North America, Des Moines, IA).
liver glycogen by 380% over values observed in a normal
fed state. Following a 22 h fast, liver glycogen concentrations in chicks were higher after 6 h compared with
2 h of refeeding (Lepkovsky et al., 1960). In the present
study, birds were refed for 30 min and then feed was removed for a period of time prior to harvest. The relative
short refeeding period employed in this study probably
was not adequate to maximize liver glycogen levels following fasting. Dietary Cr did not affect liver glycogen
following refeeding (Table 10). Lack of an effect of Cr on
liver glycogen levels in refed birds may have been due
to the short refeeding period. In turkey poults refed
for 24 h, following a 48 h fast, Cr supplementation increased liver glycogen concentrations (Rosebrough and
Steele, 1981).
In agreement with a previous study (Edwards et al.,
1999), muscle glycogen concentrations were not affected
by fasting or refeeding following fasting (Table 8). The
pooled analysis across all physiological states indicated
an overall effect of Cr (P = 0.04) on muscle glycogen.
When analyzed by physiological state Cr supplementation to the control diet did not affect muscle glycogen concentrations in fed or fasted chicks (Table 10).
Control chicks tended (P = 0.07) to have lower muscle
glycogen levels than those supplemented with Cr following refeeding. Previous studies in rats (Campbell et al.,
1989), humans (Volek et al., 2006), and sheep (Gardner
et al., 1998) have shown no effect of dietary chromium
on muscle glycogen concentrations. Sampling block did
not affect liver or muscle glycogen concentrations.
CONCLUSION
Birds supplemented with Cr, from Cr propionate, had
lower plasma NEFA concentrations than control birds
when refed following a 22 h fast. The lower circulating NEFA concentrations are consistent with increased
lipogenesis and decreased lipolysis in Cr-supplemented
birds and suggests that Cr enhanced insulin sensitivity in liver and adipose tissue. Plasma NEFA concentrations did not differ among birds supplemented with
0.2, 0.4, and 0.6 mg Cr/kg diet. Based on these results,
and under the conditions of the present study, supplementation of a diet containing 0.43 to 0.45 mg Cr/kg
with 0.2 mg Cr/kg, from Cr propionate, was adequate
to maximize insulin sensitivity.
ACKNOWLEDGEMENTS
The authors acknowledge Jill Hyda and Matt Newhouse (Kemin AgriFoods North Americas, Inc.) for
their support in monitoring this study and associated
data with the use of good laboratory practices. Appreciation is also expressed to Ilana Barasch, Jessica Nixon,
and Vickie Hedgpeth for assistance in animal sampling.
This research was partially supported by a grant from
Kemin Agrifoods North America, Inc.
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