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Am J Physiol Heart Circ Physiol 306: H373–H381, 2014.
First published November 22, 2013; doi:10.1152/ajpheart.00411.2013.
Normal cardiac function in mice with supraphysiological cardiac
creatine levels
Lucia Santacruz,1* Alejandro Hernandez,1* Jeffrey Nienaber,1* Rajashree Mishra,1 Miguel Pinilla,1
James Burchette,3 Lan Mao,2 Howard A. Rockman,2 and Danny O. Jacobs1
1
Department of Surgery, Duke University Medical Center, Durham, North Carolina; 2Department of Medicine, Duke
University Medical Center, Durham, North Carolina; and 3Department of Pathology, Duke University Medical Center,
Durham, North Carolina
Submitted 16 May 2013; accepted in final form 15 November 2013
cardiac failure; creatine transporter; energy metabolism; phosphocreatins
A HALLMARK OF CARDIAC FAILURE is a marked disturbance of
energy metabolism. In this condition, the manner in which
ATP is buffered and transferred, via the creatine kinase (CK)
system, from the mitochondria to utilization sites, is profoundly altered (26). In the early stages of heart failure,
myocardial creatine (Cr) and phosphocreatine (PCr) are reduced by as much as 70%, whereas a slow and progressive
ATP loss of up to 30 – 40% is observed at the latter stages of
the disease (4, 28, 39, 45). Moreover, the PCr-to-ATP ratio
* L. Santacruz, A. Hernandez, and J. Nienaber contributed equally to this
work.
Address for reprint requests and other correspondence: L. Santacruz, Department of Biochemistry and Molecular Biology, University of Texas, Medical Branch, Galveston, TX 77555 (e-mail: [email protected]).
http://www.ajpheart.org
decreases, and this decrease is a better predictor of overall and
cardiovascular-related mortality than traditional indexes such
as ejection fraction or New York Heart Association symptomatology (27). These deficits in high-energy metabolites are
accompanied by decreases in the activity and expression levels
of the CKs in the myocardium (22, 38). Recent reports elegantly illustrated that increased CK function has a protective
effect in murine models of global cardiac ischemia and hypertension-induced heart failure (1, 14). Cardiomyocytes are not
capable of Cr synthesis and depend on transport from the
extracellular environment by the creatine transporter (CrT) to
maintain an adequate intracellular Cr pool. CrT function is
reduced in the failing heart (29, 43), and, despite recent
progress in our understanding of cellular energetics and the
relevance of Cr and PCr in cardiac muscle, a clear understanding
of how CrT is regulated in normal and failing hearts is lacking.
We have recently shown that, in cardiomyocytes in culture, its
function is regulated by substrate availability, AMP-activated
protein kinase (AMPK) (8), and protein kinase C (9).
Given the pivotal role of Cr as a component of a spatial/
temporal energy shuttle that sustains ATP levels at sites of
energy consumption, it has been hypothesized that an increase
in myocellular Cr content would be therapeutically valuable in
the setting of acute or chronic cardiac injury. Attempts to
increase cardiac Cr content and ameliorate cardiac and energetic disarray by oral supplementation in a rat heart failure
model have proven unsuccessful (16). On the other hand,
depletion of Cr by chronic pharmacological Cr transport or
synthesis inhibition had an adverse effect on cardiac function
(19, 44). Recent characterization of physiologically relevant
regulation of cardiac Cr transport explains these earlier findings; in cardiomyocytes, Cr transport decreases quickly and
very significantly in response to increases in extracellular Cr
within the time frame and concentrations observed during oral
supplementation with Cr (8). To circumvent these limitations
and to determine if cardiac Cr content elevations would be
beneficial to stressed out cardiac muscle, transgenic mice that
overexpress the rabbit CrT were prepared. In these animals,
myocardial Cr content was augmented, but these transgenic
animals also developed heart failure spontaneously (46). It was
proposed that the increase in intracellular Cr led to increased
free ADP concentrations that in turn lowered the free energy
change of ATP hydrolysis (⌬GATP) required to drive ATP
production. However, whether these changes represented an
adaptive or causal event of cardiac failure is unknown (34, 46).
Further analysis showed that CrT overexpression, resulting in a Cr
increase below twofold that of the baseline, was well tolerated.
0363-6135/14 Copyright © 2014 the American Physiological Society
H373
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Santacruz L, Hernandez A, Nienaber J, Mishra R, Pinilla M,
Burchette J, Mao L, Rockman HA, Jacobs DO. Normal cardiac
function in mice with supraphysiological cardiac creatine levels. Am
J Physiol Heart Circ Physiol 306: H373–H381, 2014. First published
November 22, 2013; doi:10.1152/ajpheart.00411.2013.—Creatine
and phosphocreatine levels are decreased in heart failure, and reductions in myocellular phosphocreatine levels predict the severity of the
disease and portend adverse outcomes. Previous studies of transgenic
mouse models with increased creatine content higher than two times
baseline showed the development of heart failure and shortened
lifespan. Given phosphocreatine’s role in buffering ATP content, we
tested the hypothesis whether elevated cardiac creatine content would
alter cardiac function under normal physiological conditions. Here,
we report the creation of transgenic mice that overexpress the human
creatine transporter (CrT) in cardiac muscle under the control of the
␣-myosin heavy chain promoter. Cardiac transgene expression was
quantified by qRT-PCR, and human CrT protein expression was
documented on Western blots and immunohistochemistry using a
specific anti-CrT antibody. High-energy phosphate metabolites and
cardiac function were measured in transgenic animals and compared
with age-matched, wild-type controls. Adult transgenic animals
showed increases of 5.7- and 4.7-fold in the content of creatine and
free ADP, respectively. Phosphocreatine and ATP levels were two
times as high in young transgenic animals but declined to control
levels by the time the animals reached 8 wk of age. Transgenic mice
appeared to be healthy and had normal life spans. Cardiac morphometry, conscious echocardiography, and pressure-volume loop studies
demonstrated mild hypertrophy but normal function. Based on our
characterization of the human CrT protein expression, creatine and
phosphocreatine content, and cardiac morphometry and function,
these transgenic mice provide an in vivo model for examining the
therapeutic value of elevated creatine content for cardiac pathologies.
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SPECIFIC ELEVATIONS IN CARDIAC CREATINE ARE NOT TOXIC
Moreover, such an increase was clearly demonstrated to be protective in the setting of acute myocardial infarction (21).
Recently, the importance of Cr as a spatial and temporal
energy buffer in cardiomyocytes has been questioned. Knockout mice lacking the enzyme (guanidine acetate N-methyltransferase) required for the Cr biosynthesis had normal cardiac
function and, furthermore, did not appear to have worsened
outcomes following myocardial infraction (20). It is not clear how
these findings can be reconciled with reports demonstrating the
deleterious effects of Cr depletion to cardiac muscle (44).
Here, we report the generation of a transgenic mouse line
where cardiac specific overexpression of the human CrT protein resulted in a 5.7-fold increase in Cr content but with
normal ventricular function and lifespan.
Construction of human CrT transgenic mice. Human CrT-cDNA
was a kind gift from Dr. Marc Caron (Department of Cell Biology,
Duke University). The Flag epitope was introduced to the 5=-end of
the cDNA. The cDNA encoding the tagged CrT protein was subcloned into pCDNA3.0, yielding a construct where expression was
under the control of the ␣-myosin heavy chain (␣-MHC) promoter
(10). This promoter is active in both atrial and ventricular cardiomyocytes (35, 40). The resulting construct integrity was verified by
expression and function studies in Griptite cells (data not shown; Life
Technologies, Carlsbad CA). The Flag-CrT protein functional properties were similar to those obtained from the untagged CrT. Transgenic animals were generated at Duke University’s Transgenic Mouse
Facility. The above-mentioned construct was microinjected in the
pronucleus of fertilized FVB oocytes. The oocytes were implanted in
the oviducts of pseudopregnant Swiss-Webster female mice. Pups
were screened for transgene insertion by polymerase chain reactions
from DNA isolated from their tails and toes, using the 5=-aagaggcagggaagtggtgg-3= as the forward primer and the 5=-ccccagacagcctcaagact-3= as the reverse primer. Adult male mice bearing the transgene were bred with FVB mice (NCI-Frederick, Frederick, MD).
Animals were handled according to the approved protocols and
animal welfare regulations of the Institutional Review Board at Duke
University Medical Center.
Transthoracic echocardiography. Echocardiography was performed on conscious mice using either an HDI 5000 echocardiograph
(Phillips) or a Vevo 770 high-resolution imaging system (VisualSonics), as previously described by Tanaka et al. (42).
In vivo pressure-volume analysis in anesthetized mice. Mice were
anesthetized with ketamine (100 mg/kg) and xylazine (2.5 mg/kg),
and the left ventricle was cannulated with a 1.4-French conductance
catheter (Millar, Houston, TX) as previously described (32). Data
were analyzed with PVAN software (versions 3.3 and 3.6; Millar).
Immunoblotting. Mice were sedated with an intraperitoneal injection of ketamine (100 mg/kg) and xylazine (2.5 mg/kg). The thoracic
cavity was opened, and the heart was removed, rinsed in saline,
separated into the four chambers, weighed, and then flash-frozen in
liquid nitrogen in ⬃20 s. The heart was subsequently stored at ⫺80°C.
The tissue was homogenized over ice in 150 mM NaCl, 50 mM Tris,
pH 8.0, 5 mM EDTA, 1% Nonidet P-40, and 0.5% deoxycholate
supplemented with protease inhibitors (Complete Mini protease inhibitors; Roche, Indianapolis, IN) and phosphatase inhibitors (in mM:
5 NaF, 1 phenylmethylsulfonyl fluoride, 2.5 Na2P2O7, 50 ␤-glycerol,
and 1 Na3VO3; Sigma, St. Louis, MO) using an electrical tissue
homogenizer. Homogenates were centrifuged at 4°C and 100,000 g
for 45 min. The protein concentration in the supernatant was determined using the bicinchoninic acid assay (Pierce Biotechnology,
Rockford, IL), and equal amounts of protein (5.5 ␮g/lane) isolated
from wild-type (WT) or creatine transporter transgenic (CrT-Tg)
hearts were subjected to electrophoresis on a 4 –12% Novex Tris-Bis
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METHODS
precast gradient gel using MES buffer (Invitrogen, Carlsbad, CA) and
Western blot analysis, as described (7).
Immunohistochemistry. Formalin-fixed ventricles were embedded
in paraffin. Four-micrometer sections were cut from the tissue blocks,
placed on positive charged slides, allowed to air dry, and then heated
in a 65°C oven for 30 min. After removing the paraffin with xylene
and clearing with alcohol, the slides were placed in hydrogen peroxide
and methanol to quench endogenous peroxidase activity. Sections
were hydrated and washed in deionized water. It was determined
during antibody optimization that proteinase K (concentrate diluted
0.05 ml in 1.0 ml of 0.05 M Tris, pH 7.5; Dako, Carpinteria, CA) was
the proteolytic enzyme of choice. Comparative tissue pretreatment
studies were performed using heat-induced epitope retrieval. The
tissue sections were digested for 5 min in the working proteinase K
solution and then rinsed in deionized water and placed in Trisbuffered saline (TBS), pH 7.5. Primary anti-CrT rat monoclonal
antibody 4B9, (7) at a 1:500 dilution, was applied and incubated for
1 h at room temperature. Following a rinse and wash with TBS, the
bound primary antibody was linked with biotinylated goat anti-rat IgG
(H&L specific, 5 ␮g/ml; Vector Laboratories, Burlingame, CA). The
formed immune complex was further amplified and labeled with
horseradish peroxidase conjugated to the avidin biotin complex (ABC
Elite; Vector Laboratories). Diaminobenzidine was used to visualize
the bound 4B9 antibody. The slides were then washed with tap water.
Hematoxylin counterstain was applied, followed by dehydration with
absolute alcohol. Finally, the slides were cleared with xylene and
cover slipped with a permanent mounting media. For quantification of
immunostained cardiomyocytes, sections were analyzed using darkfield and phase-contrast imaging modes. Images were captured and
postprocessed using a modified version of ImageJ (National Institutes
of Health, Bethesda, MD). Sample edges were discarded to avoid
counting errors resulting from mechanical deformation of the sectioned tissue. Segmentation histograms were computed, and all segments within two SDs (total area) of the training set (hand selected)
were included in the analysis. A total of 1,935 valid segments were
selected in this manner, of which 82 corresponded to fully stained
segments (as computed from the average pixel color value). Segmentation across unstained sections presented a challenge in some areas of
the sample; thus, an average area calculation was performed as well.
The ratio of stained/unstained area (0.047) was very similar to the
result previously obtained counting single segments (0.042). The
slight increase obtained using the area comparison can be explained
because of the presence of very small segments not accounted for in
the segmentation procedure.
Real-time RT-PCR. Total RNA was extracted from left ventricle
tissue using an Aurum Total RNA Fatty and Fibrous Kit (Bio-Rad,
Hercules, CA), and genomic DNA was eliminated by DNase I
digestion. Equal RNA concentrations were used to perform a two-step
RT-PCR using oligo(dT), random hexamer primers, and the IScript
cDNA Synthesis Kit (Bio-Rad). After cDNA was obtained, qRT-PCR
was performed to determine the abundance of mRNA encoding the
CrT transgene or the native CrT relative to ␤-actin mRNA. For the
quantification of the CrT transgene, the following primers were used:
forward primer: 5=-ctgttgctgcttggtctc-3= and reverse primer: 5=-ttggaaacggaagtagtagg-3=. For ␤-actin mRNA, the primers used were:
forward primer: 5=-gacaggatgcagaaggagattact-3= and reverse primer:
5=-tgatccacatctgctggaaggt-3=, as previously described (22, 33). qRTPCR was done using iQ SYBR Green Supermix (Bio-Rad) and an MX
3005P thermal cycler (Strategene, Santa Clara, CA). Reactions were
performed in triplicate and cycle threshold values were analyzed as
reported by Pfaffl (33). Efficiencies were calculated using serial
dilutions generated from CrT transgene plasmid and WT mice RNA,
respectively.
Quantification of ATP, ADP, Cr, and PCr. Beating hearts were
clamped while still in the chest using liquid N2 precooled clamps. The
frozen heart “wafers” were stored at ⫺80°C until extract preparation.
Samples were submerged in liquid N2, weighed, and ground to a fine
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SPECIFIC ELEVATIONS IN CARDIAC CREATINE ARE NOT TOXIC
RESULTS
Phenotype and transgene expression. Expression of the
transgene in CrT-Tg animals was characterized by qRT-PCR
(Table 1), Western blotting (Fig. 1A), and immunohistochemical analysis (Fig. 1B). The mRNA encoding the human CrT
and native CrT protein was quantified in 2-, 4-, and 8-wk-old
animals. The mRNA encoding transgene (human CrT) was
⬃60 times more abundant than that encoding the native
(mouse) CrT protein (Table 1). We observed a decrease in the
mRNA encoding for both the native and human CrT in 4-wkold CrT-Tg, but both rebounded in 8-wk-old Cr-Tg animals,
resulting in 8-wk-old mice having the same expression level
for the transgene message as 2-wk-old animals. In WT animals,
the level of mRNA for native CrT increased as the animals
matured, a trend also observed in transgenic animals (Table 1).
The presence of the human CrT protein was also verified by
probing Western blots of cardiac tissue homogenates from
6-mo-old CrT-Tg and WT animals with an anti-CrT antibody
that preferentially recognized the human isoform of the protein
(7). A band of ⬃55 kDa was observed only in the solubilized
homogenates from CrT-Tg hearts (Fig. 1A). Immunohistochemistry was used to further document the expression of the
protein encoded by the transgene in cardiac muscle sections.
Interestingly, not every cardiomyocyte was stained, and quantification of these cells in the ventricle sections used for
immunohistochemistry indicated that 4% of the cardiomyocytes were immunopositive. This “mottled” pattern was observed in hearts from both male and female transgenic animals,
thereby excluding integration of the transgene in the X-chro-
mosome. A similar nonhomogenous expression pattern was
previously reported for transgenes under the control of the
␣-MHC promoter (31). In the cells that were stained by the
anti-CrT antibody, the signal concentrated at the cell membrane and had a striated pattern. This staining was not observed
in WT animals (Fig. 1B).
Cardiac energy metabolite measurements. Cr, PCr, ATP,
and ADP were quantified in flash-frozen cardiac tissue extracts
from 2-, 4-, and 8-wk-old CrT-Tg animals and age-matched
WT controls by HPLC, and free ADP was calculated as
described in METHODS. CrT-Tg animals had significantly higher
myocellular Cr levels (Fig. 2A). By the time animals reached
adulthood (8 wk), cardiac Cr was 5.7-fold higher in 8-wk-old
CrT-Tg mice than in WT animals. PCr content was also
significantly elevated in 2- and 4-wk-old CrT-Tg mice, but, by
adulthood, had decreased to levels observed in WT agematched animals (Fig. 2B). Similarly, ATP concentrations
were also significantly higher in 2- and 4-wk-old CrT-Tg mice.
However, ATP levels were lower than those of age-matched
controls by week 8 (Fig. 2C). Total ADP (tADP) levels were
significantly reduced in 8-wk-old CrT-Tg animals compared
with WT animals. There were no significant differences in
tADP content in younger animals (Fig. 2D). Free ADP content
was determined as described in METHODS and found to be
significantly elevated at all ages in CrT-Tg hearts compared
with WT hearts (Fig. 2E). The resulting PCr-to-ATP ratio was
significantly elevated in 4- and 8-wk-old CrT-Tg mice compared with WT age-matched controls (Fig. 2F), whereas PCrto-Cr ratios were reduced in CrT-Tg hearts at all ages (Fig.
2G), likely reflecting the increased size of the Cr pool secondary to augmented Cr transport. The ATP-to-tADP ratio (Fig.
2H) was significantly elevated in CrT-Tg 2- and 4-wk-old
animals compared with the respective age-matched WT animals. This difference was not observed in 8-wk-old animals.
Cardiac morphometry and function. The morphometry and
function of CrT-Tg hearts was analyzed throughout development and into maturity and compared with WT age-matched
animals (Tables 2 and 3). Significant increases were observed
in the right atrium-to-tibial length ratio in 2-, 4-, and 8-wk- and
9-mo-old CrT-Tg animals and the left atrium-to-tibial length
ratio of 4- and 8-wk- and 9-mo-old CrT-Tg animals. Although
the LV/body weight ratios were not significantly different, the
LV/TL ratios were slightly but significantly greater at all ages
among CrT-Tg compared with WT animals. Lung weight was
significantly less among CrT-Tg at 4 and 8 wk. Echocardiography showed no differences in fractional shortening or left
ventricular chamber diameters at any time point and significantly increased posterior wall thickness at 4 wk and 9 mo
(Table 2). Taken together, these data indicate mild left ven-
Table 1. Transcription of CrT encoding mRNA in CrT-Tg and wild-type mice
2 Weeks (n ⫽ 6)
Native CrT mRNA
Human CrT mRNA
4 Weeks (n ⫽ 5)
8 Weeks (n ⫽ 6)
CrT-Tg
WT
CrT-Tg
WT
CrT-Tg
WT
0.0073 ⫾ 0.001
0.56 ⫾ 0.17
0.0050 ⫾ 0.0003
NA
0.0037 ⫾ 0.0008
0.22 ⫾ 0.06
0.0108 ⫾ 0.0013
NA
0.014 ⫾ 0.001
0.63 ⫾ 0.11
0.0162 ⫾ 0.0017
NA
Values are means ⫾ SE; n, no. of animals. Real-time quantification of mRNA encoding the native (mouse) and transgene (human) forms of the creatine
transporter (CrT) protein in both creatine transporter transgenic (CrT-Tg) and wild-type (WT) age-matched control animals. The abundance of the respective
mRNAs was determined in 2-, 4-, and 8-wk-old animals and was normalized to ␤-actin and expressed in arbitrary units based on the cycle threshold (Ct) values,
as described in METHODS. NA, not applicable.
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powder using disposable individual tissue grinders. Tissue extracts
were prepared and subjected to HPLC analysis following the method
described by Wiseman et al. (47), assuming an intracellular volume of
0.5 ml/g wet wt. ATP, Cr, and PCr were measured by HPLC, and free
ADP levels were calculated based on the CK equilibrium equation:
[ADP] ⫽ [ATP][Cr]/[PCr] ⫻ KCK where square brackets denote
concentration. The value for the creatine kinase equilibrium constant
共KCK兲 depends on the pH and free Mg2⫹ measurements. Thus, we
used the values determined by phosphorus nuclear magnetic resonance spectroscopy in murine hearts as reported by Himmelreich and
Dobson (free Mg2⫹ ⫽ 0.4 mM, pH ⫽ 7.32) (15). The KCK at 38°C
was as reported by Golding et al. (13). Metabolite ratios were
calculated using only data from heart extracts where values for both
metabolites (e.g., PCr and ATP) were obtained.
Statistical analyses. Data are reported as means ⫾ SE and were
analyzed using STATISTICA 6.1 (StatSoft, 1984 –2003) with one- or
two-way ANOVA for multiple groups of independent samples, followed by post hoc, pairwise comparisons. A probability value of
ⱕ0.05 was considered significant. Most group comparisons had a 1:1
proportion of male to female animals. Kaplan-Meyer survival analysis
was performed using Graphpad Prism (GraphPad, La Jolla, CA).
H376
SPECIFIC ELEVATIONS IN CARDIAC CREATINE ARE NOT TOXIC
A
CrT-Tg
WT
75 kDa
50 kDa
B
WT
100 µm
CrT-Tg
100 µm
tricular hypertrophy and atrial enlargement without echocardiographic evidence of chamber enlargement of contractile
dysfunction among CrT-Tg animals. Left ventricular function
was further evaluated in vivo by pressure-volume loop analysis. At both 8 wk and 9 mo (Table 3), there were no significant
differences in systolic or diastolic function between CrT-Tg
and WT mice. Kaplan-Mayer plots of WT and CrT-Tg animals
overlapped, indicating that lifespans and mortality rates were
not different from those of controls animals (data not shown).
DISCUSSION
We report the creation of a transgenic mouse that expresses
the human Cr transporter in the heart with normal cardiac
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Fig. 1. Human creatine transporter (hCrT) is expressed
in creatine transporter transgenic (CrT-Tg) cardiac muscle.
A: Western blot of cardiac tissue homogenates from adult
transgenic or wild-type (WT) mice. In CrT-Tg homogenates, a
band of ⬃55 kDa corresponding to the hCrT protein was
detected using a specific rat monoclonal antibody against the
human CrT protein isoform. B: cardiac ventricle tissue sections
from 8-wk-old CrT-Tg animals or WT age-matched controls
were prepared for histological analysis as described in METHODS. Staining was observed only in CrT-Tg animals (bottom),
and localized to the cell surface in a striated pattern.
H377
SPECIFIC ELEVATIONS IN CARDIAC CREATINE ARE NOT TOXIC
40
9
*
30
25
*
20
8
10
15
10
5
0
4
Total ADP (mM)
#
20
6
9
7
10
Wild type
140
10
10
10
*
12
2
9
1.5
120
4 weeks
Wild type
F
*
*
*
6
10
60
12
CrT-Tg
5
6
7
6
6
4
3.00
ATP/ADP (ratio)
8
5
6
4
3
6
*
*
10
9
6
5
2
1
4 weeks
8 weeks
2 weeks
4 weeks
8 weeks
4.00
3.50
Wild Type
10
Wild type
0
2 weeks
H
8 weeks
CrT-Tg
7
9
80
6
4 weeks
6
100
8 weeks
9
12
2 weeks
8 weeks
0
4 weeks
5
4
#*
0
20
2 weeks
10
1
40
0
10
6
2
160
10
10
3
5
CrT-Tg
1
PCr/Cr (ratio)
6
15
Wild type
10
7
25
180
0.5
G
10
2 weeks
E
CrT-Tg
3
2.5
*
30
*
*
*
CrT-Tg
Wild type
10
10
2.50
2.00
10
10
9
12
1.50
1.00
2
5
*
*
6
10
0.50
*
9
0
2 weeks
4 weeks
8 weeks
0.00
2 weeks
4 weeks
8 weeks
Fig. 2. High-energy metabolite analysis. Hearts from control or CrT-Tg mice were harvested, and the content of creatine (Cr), phosphocreatine (PCr), ATP, and
free ADP was determined by HPLC as described in METHODS. The number within each bar indicates the number of hearts analyzed. A: CrT-Tg mice hearts had
significantly higher Cr content compared with WT mice. B: PCr levels were significantly increased in CrT-Tg levels in 2- and 4-wk-old animals (2.3- to 2.5-fold
vs. WT controls). By 8 wk of age, the content of PCr in CrT-Tg mice was the same as WT controls. C: ATP levels were significantly elevated in CrT-Tg at 2
and 4 wk of age but fell below those measured in WT animals in 8-wk-old CrT-Tg hearts. D: total ADP content was significantly elevated in 8-wk-old WT
animals. Two- and 4-wk-old animals had similar ADP content, irrespective of transgene expression. E: free ADP content (calculated as described in METHODS)
was elevated in CrT-Tg animals at all time points. F: the PCr-to-ATP ratio was significantly elevated in CrT-Tg animals at 4 and 8 wk of age. G: CrT-Tg animals
had significantly greater PCr-to-Cr ratios than age-matched WT controls. H: the ATP-to-ADP ratio (calculated using the measured total ADP content) was
significantly higher in 2- and 4-wk-old CrT-Tg animals compared with WT animals. This difference was not observed in 8-wk-old animals. *P ⬍ 0.05 vs. WT,
age-matched control, 2-way ANOVA, Fisher’s least significant difference (LSD). #P ⬍ 0.05 vs. same line at 2 wk, 2-way ANOVA, Fisher’s LSD.
function and survival. Cardiac Cr content in CrT-Tg animals
was nearly six times higher than that of WT mice as demonstrated by HPLC analysis. In addition, CrT protein expression
was documented by Western blotting and immunohistochemical analysis of cardiac muscle sections (Fig. 1B), demonstrating for the first time that functional CrT protein is found at the
cell membrane. Approximately 4% of the cardiomyocytes in
the sections subjected to immunohistochemical analysis had
robust staining with the anti-CrT antibody. Nonhomogenous
cardiac expression of a transgene driven by the ␣-MHC promoter has been previously reported (31).
No functional ventricular cardiac abnormalities were observed on echocardiographic or pressure-volume studies, and
the structural differences observed in morphometric studies
indicate a mild left ventricular hypertrophy with no deleterious
effects on cardiac function (Tables 2 and 3). Our results differ
significantly from those initially described by Wallis et al.
where mice with two- to fourfold increases in intracellular Cr
concentrations developed cardiac dysfunction spontaneously
(21, 34, 46). Moreover, in these animals, the severity of
dysfunction was positively correlated with intracellular Cr
concentrations. The authors hypothesized that the increase in
total Cr led to increased free ADP concentration, which in turn
caused the free energy change of ATP hydrolysis (⌬GATP) to
decrease. In this scenario, and in conjunction with a CK system
that remained at the same functional level as in WT animals,
the Cr shuttle’s main function (keeping ADP levels low and
⌬GATP high) would be impaired. The authors also reported
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3.5
8
35
8 weeks
free ADP (µM)
D
4 weeks
*
9
*
0
2 weeks
CrT-Tg
10
Wild type
5
10
10
9
C
CrT-Tg
ATP (mM)
# *
Wild type
35
Creatine (mM)
45
PCr/ATP (ratio)
40
B
CrT-Tg
Phosphocreatine (mM)
A 45
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SPECIFIC ELEVATIONS IN CARDIAC CREATINE ARE NOT TOXIC
Table 2. Cardiac morphometry and function of CrT-Tg mice
2 Weeks
8 Weeks
9 Months
CrT-Tg (n ⫽ 10)
WT (n ⫽ 10)
CrT-Tg (n ⫽ 10)
WT (n ⫽ 10)
CrT-Tg (n ⫽ 8)
WT (n ⫽ 10)
CrT-Tg (n ⫽ 24)
WT (n ⫽ 26)
7.20 ⫾ 0.27
10.70 ⫾ 0.20
97.37 ⫾ 2.50
8.47 ⫾ 0.38
26.16 ⫾ 0.66*
3.66 ⫾ 0.09
2.45 ⫾ 0.03*
0.77 ⫾ 0.05
0.14 ⫾ 0.01
0.22 ⫾ 0.01*
547 ⫾ 21
0.48 ⫾ 0.04
0.53 ⫾ 0.04
65.82 ⫾ 3.48
2.41 ⫾ 0.06
0.82 ⫾ 0.08
7.05 ⫾ 0.24
10.63 ⫾ 0.12
98.17 ⫾ 2.34
8.23 ⫾ 0.52
22.41 ⫾ 0.55
3.74 ⫾ 0.12
2.11 ⫾ 0.06
0.80 ⫾ 0.04
0.13 ⫾ 0.01
0.15 ⫾ 0.01
535 ⫾ 14
0.54 ⫾ 0.05
0.51 ⫾ 0.04
67.58 ⫾ 3.11
2.48 ⫾ 0.07
0.81 ⫾ 0.09
16.95 ⫾ 0.61
15.37 ⫾ 0.22
118.56 ⫾ 3.42*
8.47 ⫾ 0.38
63.28 ⫾ 1.64*
3.76 ⫾ 0.10
4.11 ⫾ 0.06*
1.14 ⫾ 0.05
0.27 ⫾ 0.02*
0.41 ⫾ 0.06*
696 ⫾ 7.2
0.76 ⫾ 0.05
0.87 ⫾ 0.06*
65.57 ⫾ 3.39
2.93 ⫾ 0.06
1.02 ⫾ 0.11
17.63 ⫾ 0.50
15.32 ⫾ 0.09
125.00 ⫾ 3.32
8.23 ⫾ 0.52
55.04 ⫾ 1.14
3.13 ⫾ 0.05
3.59 ⫾ 0.06
1.08 ⫾ 0.08
0.21 ⫾ 0.01
0.21 ⫾ 0.01
691 ⫾ 6.5
0.66 ⫾ 0.06
0.69 ⫾ 0.03
64.23 ⫾ 1.36
3.03 ⫾ 0.10
1.09 ⫾ 0.05
24.16 ⫾ 1.34
17.54 ⫾ 0.22
125.51 ⫾ 4.40*
22.31 ⫾ 1.49
79.53 ⫾ 4.33*
3.30 ⫾ 0.09
4.53 ⫾ 0.24*
1.27 ⫾ 0.08
0.30 ⫾ 0.02*
0.59 ⫾ 0.06*
619 ⫾ 10.2
0.95 ⫾ 0.03
0.85 ⫾ 0.05
45.33 ⫾ 1.33
3.13 ⫾ 0.10
1.71 ⫾ 0.08
24.09 ⫾ 1.06
17.56 ⫾ 0.12
137.59 ⫾ 3.76
20.45 ⫾ 1.48
73.97 ⫾ 1.86
3.10 ⫾ 0.08
4.21 ⫾ 0.09
1.16 ⫾ 0.08
0.24 ⫾ 0.02
0.25 ⫾ 0.01
673 ⫾ 8.3
0.89 ⫾ 0.04
0.84 ⫾ 0.02
48.12 ⫾ 1.59
3.06 ⫾ 0.05
1.59 ⫾ 0.06
40.5 ⫾ 1.82
18.84 ⫾ 0.09
157.66 ⫾ 4.30
28.86 ⫾ 1.55
114.11 ⫾ 4.13*
2.86 ⫾ 0.08
6.05 ⫾ 0.20*
1.53 ⫾ 0.08*
0.49 ⫾ 0.03*
0.46 ⫾ 0.04*
645 ⫾ 7.1
1.07 ⫾ 0.03
1.15 ⫾ 0.04*
47.37 ⫾ 1.39
3.67 ⫾ 0.07
1.93 ⫾ 0.06
36.07 ⫾ 1.36
18.61 ⫾ 0.06
162.12 ⫾ 4.14
24.71 ⫾ 0.93
104.90 ⫾ 3.79
2.95 ⫾ 0.10
5.63 ⫾ 0.20
1.33 ⫾ 0.05
0.31 ⫾ 0.02
0.27 ⫾ 0.02
653 ⫾ 10.1
1.10 ⫾ 0.04
0.99 ⫾ 0.04
51.03 ⫾ 1.50
3.60 ⫾ 0.06
1.77 ⫾ 0.07
Values are means ⫾ SE; n, no. of animals. BW, body wt; HR, heart rate; SW, septal wall thickness; PW, posterior wall thickness; LVDd, left ventricle
dimension (diastole); LVDs, left ventricle dimension (systole). The mass of the left ventricle (LV), left atrium (LA), right ventricle (RV), or right atrium (RA)
was normalized to the corresponding animal’s tibial length (TL) and compared between age-matched CrT-Tg and WT control animals. The LV-to-TL and
RA-to-TL ratios of CrT-Tg animals were significantly greater at all ages compared with WT controls, as were the LA-to-TL ratios of 4- and 8-wk- and 9-mo-old
CrT-Tg animals. The percent fractional shortening (%FS), left ventricular chamber dimensions, and septal/posterior wall thicknesses were determined in
conscious 2-, 4-, and 8-wk-old and 9-mo-old mice as described in METHODS. Posterior wall thickness was significantly greater at 4-wk and 9-mo among CrT-Tg.
No other echocardiographic differences were detected between CrT-Tg mice and age-matched WT control animals. *Significant difference with age-matched WT
controls, P ⬍ 0.05 (ANOVA, Tukey’s honest significant difference).
diminished ␣-enolase activity and reduced flux through the
glycolytic pathway. However, whether these changes represented an adaptive or a causal event of cardiac failure was
unknown (34). Upon further screening of this transgenic animal model, it was determined that very moderate increases in
Cr transport capacity, up to a twofold increase in cardiac Cr
content, were well tolerated. More importantly, the authors
demonstrated that these increases afforded protection from
ischemic insults to the cardiac muscle (21).
There are several possible explanations for the discrepancies
noted between the transgenic line studied here and those
reported in the studies by Wallis and Phillips. First, mice of
different genetic backgrounds may react differently to the same
cardiac stressor (3, 6, 36, 41). In the studies by Wallis and
Phillips, C57BL/6 mice genetic backgrounds were used,
whereas FVB mice were used in our study. C57BL/6 mice
appear to be more susceptible to developing cardiomyopathy
and have increased mortality rates following transverse aortic
constriction compared with other strains (3). FVB mice tend to
develop myocardial hypertrophy without severe contractile
alterations after the same intervention (10, 12). Thus it is
plausible that differences in animal strains may be involved in
the adaptation of the heart to mechanical as well as to biochemical stress factors. In an effort to address this specific
issue, we have begun backcrossing the CrT-Tg mice into the
C57/Bl6 background. At this time, we have bred the sixth
Table 3. Cardiac function of adult and aged CrT-Tg animals: pressure-volume loop measurement
8 Weeks
WT (n ⫽ 10)
Systolic function
Heart rate, beats/min
End-systolic volume, ␮l
End-diastolic volume, ␮l
Maximum SBP, mmHg
End-systolic pressure, mmHg
dP/dtmax, mmHg/s
Cardiac output, ␮l/min
Ejection fraction
End-systolic elastance
Diastolic Function
End-diastolic pressure, mmHg
dP/dtmin, mmHg/s
Tau (Glatz), ms
EDPVR
9 Months
CrT-Tg (n ⫽ 7)
WT (n ⫽ 8)
CrT-Tg (n ⫽ 9)
399 ⫾ 15
17.17 ⫾ 1.48
35.94 ⫾ 2.34
91.35 ⫾ 3.67
83.68 ⫾ 4.76
8,302 ⫾ 518
8.70 ⫾ 0.40
58.75 ⫾ 2.47
5.78 ⫾ 0.56
397 ⫾ 17
17.13 ⫾ 3.63
32.36 ⫾ 2.78
85.34 ⫾ 1.46
77.05 ⫾ 3.18
9,176 ⫾ 604
7.51 ⫾ 0.92
56.47 ⫾ 6.96
5.23 ⫾ 0.56
455.44 ⫾ 15.05
7.15 ⫾ 0.96
17.04 ⫾ 1.81*
86.46 ⫾ 3.45
75.01 ⫾ 4.62
8,099 ⫾ 460
5.05 ⫾ 0.50*
62.69 ⫾ 2.84
7.41 ⫾ 1.07*
399.14 ⫾ 24.37
7.74 ⫾ 0.97Ŧ
17.91 ⫾ 1.80Ŧ
81.04 ⫾ 3.09
71.13 ⫾ 3.09
8,535 ⫾ 710
4.54 ⫾ 0.21Ŧ
62.63 ⫾ 1.66
6.32 ⫾ 0.50Ŧ
7.98 ⫾ 1.20
⫺6,773 ⫾ 503
10.87 ⫾ 0.41
0.62 ⫾ 0.09
5.64 ⫾ 0.61
⫺6,116 ⫾ 310
11.57 ⫾ 1.07
0.80 ⫾ 0.18
3.74 ⫾ 0.40*
⫺6,996 ⫾ 719
10.88 ⫾ 0.79
0.97 ⫾ 0.15
4.08 ⫾ 0.56Ŧ
⫺6,000 ⫾ 319
12.01 ⫾ 0.61
0.84 ⫾ 0.14
Values are means ⫾ SE; n, no. of animals. SBP, systolic blood pressure; EDPVR, end-diastolic pressure volume relationship. Cardiac function was evaluated
by pressure volume measurements as described in METHODS in 8-wk- and 9-mo-old animals. Significant differences between the 8-wk-old animals compared with
9-mo-old animals of the same genotype are demoted by for WT (*) or CrT-Tg (Ŧ) (P ⬍ 0.05, 2-way ANOVA, nonrepeated measures and Bonferroni post hoc
test).
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Body wt, g
TL, mm
Lungs, mg
RV wt, mg
LV wt, mg
LV/BW, mg/g
LV/TL, mg/mm
RV/TL, mg/mm
LA/TL, mg/mm
RA/TL, mg/mm
HR, beats/min
SW, mm
PW, mm
FS, %
LVDd, mm
LVDs, mm
4 Weeks
SPECIFIC ELEVATIONS IN CARDIAC CREATINE ARE NOT TOXIC
activity is lowered by product accumulation (18, 24). Moreover, it was proposed that direct ATP channeling between the
mitochondria, the myofibrils, and the sarcoplasmic reticulum
might increase when the CK system becomes insufficient (18,
37). Thus, whereas both our models demonstrate an increase in
free ADP concentration, which would lower the energy available for ATPase activity required for excitation-contraction
coupling, the results of our functional studies suggest the
existence of an alternative pathway that allows the cell to
maintain and buffer ATP within normal ranges at the sites of
consumption (contractile machinery and ion pumps). This
mechanism may also be responsible for the capacity of CrT-Tg
juvenile mice capacity to maintain an increased ATP and
ATP-to-tADP ratios (at 2 and 4 wk of age) despite elevated
free ADP concentrations during maturation to adulthood,
which could explain why these mice did not develop heart
failure. Understanding how these developing mice were able to
maintain high ATP levels despite high free ADP concentrations could shed light on the mechanisms cardiomyocytes use
to buffer high-energy metabolites and respond to functional
and biochemical demands and potential therapeutic interventions to ameliorate the energy derangement observed in failing
hearts.
Previously we showed in HL-1 cells and rat neonatal cardiomyocytes that Cr transport decreases rapidly in response to
increased Cr in the extracellular environment. Similar observations were made in isolated heart preparations (5). The
molecular mechanism by which the cell reduces Cr transport in
response to increased Cr is largely unknown. However, we
have shown that, in cardiomyocytes in culture, the decrease in
Cr transport is the result of a reduction in the cell surface
content of CrT transporter protein (8). The expression of CrT
protein detected on Western blots from CrT-Tg animals appeared robust (Fig. 1, A and B). The mRNA encoding the
native transporter increased as the animals matured in both WT
and CrT-Tg mice, just as it occurs in developing rat hearts (11).
These observations suggest that the increased intracellular Cr
present in CrT-Tg animals did not exert a significant repressive
effect at the transcriptional level (Table 1) and that the chronically increased cardiac Cr content did not cause a decrease in
the content of human CrT mRNA. This observation would also
be consistent with what is known about substrate regulation of
Cr transport, i.e., increased extracellular Cr content decreases
Cr transport capacity.
A significant question remains surrounding the impact of
sustained Cr, PCr, and free ADP elevations on intracellular pH
and key energy homeostasis enzymatic complexes, such as
AMPK, acetyl-CoA carboxylase, or lactate dehydrogenase, all
of which are perturbed in heart failure (2, 11, 25). In addition
of answering these questions, the fact that not every cardiomyocyte in the CrT-Tg heart appears to express the human CrT
protein may be particularly useful in answering how elevated
Cr content is cardioprotective, because within the same organ
(heart) the effects of Cr elevations can be studied and contrasted in vivo to control cells. Another interesting question is
how the additional Cr that is brought into the cells expressing
the transgene diffuses into neighboring cells, increasing Cr
content “by proxy” and what would be the effect on the overall
metabolic state of these cells. In particular, given our observation that a small number of modified cardiomyocytes (4%) can
cause a significant elevation of cardiac Cr content, the explo-
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generation, and these animals do not display any sign of
cardiac malfunction. Second, the site of transgene insertion
may disrupt sensitive coding or regulatory gene(s) sequences
with the end result being the development of cardiac dysfunction phenotype. Notably, transgenic animals expressing green
fluorescent protein in cardiac muscle may spontaneously develop heart failure (17), raising the possibility that the site or
copy number of transgene insertion(s) may be responsible for
adverse cardiac effects unrelated to the function of the transgene itself. During the initial stages of our study, we isolated
several founders that died as pups, in addition to a transgenic
lineage that did develop heart failure. This lineage also had
higher expression of the human CrT, and the Cr content was
increased 8- to 10-fold than that measured in WT littermates
(data not shown). We did not determine the copy number or
site of insertion of the transgene for any of our lines, and thus
we cannot rule out that in these transgenic animals the deleterious effects were the result of genetic disruptions or cellular
toxicity because of the very elevated Cr content. Taken together with the findings from other CrT overexpression models, our results suggest that the heart can tolerate very significant (6-fold) and chronic increases in cardiac Cr content.
Third, although both mouse models used cardiac specific
promoters (MLC2v, Wallis et al., and ␣-MHC in our study),
they differ in their spatial-temporal expression activation profile during gestation and after birth, which could influence how
the transgene is regulated (23, 30). Last, the two transgenic
models differ in the species of the transgene inserted. In our
model we used the human isoform, whereas Wallis and colleagues used a rabbit CrT. Protein sequence analysis (ClustalO
Uniprot) shows that the human CrT (Uniprot identifier P48029)
is 98.12% identical to the mouse CrT (Uniprot identifier
Q8VBW1), whereas the rabbit CrT (Uniprot Identifier P31661)
amino acid sequence differs slightly more (96.87% identity).
These differences are very small and consist mostly of conserved amino acid changes. Thus, it seems unlikely that sequence differences could underlie the markedly different phenotypes of the two transgenic lines.
CK, a key element in the “creatine energy shuttle,” is the
conduit by which high-energy phosphates are transferred from
the sites of generation (ATP in the mitochondria) to sites of
consumption (contractile machinery and ion pumps) using PCr
as a carrier. CK function and distribution analysis indicated
that the transgenic animals generated by Wallis and colleagues
had no changes in the activity or distribution of CK isoforms
(46). Interestingly, the high-energy phosphate metabolite profile observed in our CrT-Tg mice was similar to that reported
by Wallis and Phillips. Namely, PCr and ADP levels were
elevated in adult animals, whereas ATP levels in the same age
group were below those observed in control animals. In our
transgenic model, we observed that 2- and 4-wk-old CrT-Tg
animals had elevated ATP content, which decreased to below
the levels measured in WT littermates (controls) by the time
the animals were 8 wk old. Although we did not measure the
activity or distribution of CK isotypes in CrT-Tg animals, the
similarities in the profiles of ATP, ADP, and PCr suggest that
Cr kinase enzymatic capabilities of CrT-Tg were not altered.
Previous reports had hypothesized that ADP removal from
areas surrounding cellular sites with elevated ATPase activity
(i.e., myosin ATPase and sarcoplasmic reticulum ATPase)
helps regulate energy regeneration locally, e.g., where ATPase
H379
H380
SPECIFIC ELEVATIONS IN CARDIAC CREATINE ARE NOT TOXIC
ACKNOWLEDGMENTS
6.
7.
8.
9.
10.
11.
We acknowledge Lauren Goers for technical assistance, Quique Toloza for
assistance in the preparation of this manuscript, Eric Toloza and Marcus
Darrabie for assistance with statistical analysis, and Dawn Bowles, Kumar
Pandya, and Bryan Feger for critical reading and insightful comments.
Current addresses: L. Santacruz, Dept. of Biochemistry and Molecular
Biology, University of Texas, Medical Branch, Galveston, TX 77555; A.
Hernandez, Dept. of Anesthesiology, Yale University School of Medicine,
New Haven, CT; J. Nienaber, Charles George VA Medical Center, 1100
Tunnel Rd., Asheville, NC 28805; M. Pinilla, UPMC Montefiore Hospital,
N-715, 200 Lothrop St., Pittsburgh, PA 15213; and D. O. Jacobs, School of
Medicine, University of Texas Medical Branch, 301 University Blvd., Galveston, TX 77555-0133.
12.
13.
14.
GRANTS
This work was funded by the Department of Surgery at Duke University
Medical Center and National Heart, Lung, and Blood Institute Grants HL056687 and T32-HL-07101 to H. A. Rockman, which provided funding to J.
Nienaber.
15.
16.
DISCLOSURES
No conflicts of interest, financial or otherwise, are declared by the authors.
17.
AUTHOR CONTRIBUTIONS
Author contributions: L.S., A.H., and J.N. conception and design of research; L.S., A.H., J.N., R.M., M.P., J.B., and L.M. performed experiments;
L.S., A.H., J.N., M.P., and L.M. analyzed data; L.S., A.H., J.N., M.P., and
L.M. interpreted results of experiments; L.S. prepared figures; L.S. drafted
manuscript; L.S., A.H., J.N., M.P., L.M., H.A.R., and D.O.J. edited and revised
manuscript; L.S., A.H., J.N., R.M., M.P., J.B., L.M., H.A.R., and D.O.J.
approved final version of manuscript.
18.
19.
20.
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