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Monitoring tissue development in acellular matrix-based regeneration for bladder tissue engineering: Multiexponential diffusion and
T2* for improved specificity
Hai-Ling Margaret Cheng, Yasir Loai, Walid A. Farhat
Version Post-print/accepted manuscript
Citation (published version)
Cheng HL, Loai Y, Farhat WA. Monitoring tissue development in acellular matrix‐based regeneration for bladder tissue engineering: Multiexponential diffusion and T2* for improved specificity. NMR in Biomedicine. 2012 Mar;25(3):418-26.
Publisher’s Statement This is the peer reviewed version of the following article: [Cheng HL, Loai Y, Farhat WA. Monitoring tissue development in acellular matrix‐based regeneration for bladder tissue engineering: Multiexponential diffusion and T2* for improved specificity. NMR in Biomedicine. 2012 Mar;25(3):418-26.], which has been published in final form at [10.1002/nbm.1617]. This article may be used for non-commercial purposes in accordance with Wiley Terms and Conditions for Self-Archiving.
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MRI of acellular matrix-based bladder tissue engineering
1
Monitoring tissue development in acellular matrix-based
regeneration for bladder tissue engineering:
multiexponential diffusion and T2* for improved specificity
Hai-Ling Margaret Cheng, PhDa,b*
, Yasir Loai, BScc, Walid A. Farhat, MD
c
a Department of Medical Biophysics, University of Toronto, Toronto, ON, Canada
b Physiology & Experimental Medicine, The Research Institute and Diagnostic
Imaging, The Hospital for Sick Children, Toronto, ON, Canada
c Developmental and Stem Cell Biology, The Research Institute and Division of
Urology, The Hospital for Sick Children, Toronto, ON, Canada
Short title: MRI of acellular matrix-based bladder tissue engineering
Grant sponsor: CIHR – Institute of Human Development Child and Youth Health /
SickKids Foundation (XG 08-017)
* Correspondence to:
Hai-Ling Margaret Cheng, PhD
Department of Diagnostic Imaging
The Hospital for Sick Children
555 University Avenue
Toronto, Ontario M5G 1X8, Canada
(tel) 416-813-5415 (fax) 416-813-7362
E-mail: hai-ling.cheng@sickkids.ca
MRI of acellular matrix-based bladder tissue engineering
2
ABSTRACT
Cell-seeded acellular matrices (ACMs) are a promising approach for tissue-
engineering soft tissues and organs such as the urinary bladder. The ACM contains site-
preferred structural and functional molecules, and degradation products derived from the
ACM play important roles in tissue remodeling. Regeneration proceeds along concurrent
trajectories of cell growth and matrix degradation, characterized by evolving biophysical
and biochemical properties. Assessment of tissue development through a non-invasive
imaging technique such as MRI must, therefore, be capable of distinguishing these
concurrent biophysical and biochemical changes. However, although MRI provides
exquisite sensitivity to tissue microstructure, composition, and function, specificity remains
limited. In this study, multiexponential diffusion and effective transverse relaxation time
T2* were investigated for their ability to assess cell growth and tissue composition,
respectively. Bladder ACMs prepared with and without hyaluronic acid, and ACMs seeded
with smooth muscle cells, were assessed on MRI. The slow diffusion fraction from
multiexponential diffusion analysis demonstrated the best correlation with cellularity, with
minimal influence from underlying matrix degradation. T2* measurements were sensitive
to macromolecular content, specifically, the presence of hyaluronic acid, without
confounding influence from tissue hydration. T2* also appeared sensitive to cell filling of
matrix pore space. Compared to these metrics, commonly used MRI parameters such as T1,
T2, and single diffusion coefficient were more limited in specificity. Use of T2 to measure
tissue structure and composition is limited by its large dependence on water content, and
single diffusion can only reflect the overall characteristics of the extra- and intracellular
MRI of acellular matrix-based bladder tissue engineering
3
environment. These findings are important for further development of more specific MRI
methods for monitoring regeneration in tissue engineered systems.
Keywords: quantitative (MRI) magnetic resonance imaging; multiexponential diffusion;
(T2*) effective transverse relaxation time; (ACM) acellular matrix; cellularity; (GAG)
glycosaminoglycan; tissue hydration; bladder tissue engineering
Abbreviations:
2D, two-dimensional
3D, three-dimensional
ACM, acellular matrix
dH2O, double distilled water
GAG, glycosaminoglycan
HA, hyaluronic acid
HBSS, Hank’s Balanced Salt Solution
H&E, hematoxylin and eosin
VEGF, vascular endothelial growth factor
MRI of acellular matrix-based bladder tissue engineering
4
GRAPHICAL ABSTRACT
Monitoring tissue development in acellular matrix-based regeneration for bladder tissue
engineering: multiexponential diffusion and T2* for improved specificity
Hai-Ling Margaret Cheng, PhD*, Yasir Loai, BSc, Walid A. Farhat, MD
Multiexponential diffusion MRI and T2* are investigated for monitoring acellular matrix-
based bladder regeneration. The slow diffusion fraction was shown to correlate with
cellularity in the presence of extracellular matrix changes, while T2* was sensitive to
macromolecules without influence from tissue hydration. These metrics provide improved
specificity over single T1, T2, and diffusion for assessing concurrent tissue engineering
processes.
I II III IV
0
0.1
0.2
T2* T2 T1
Higher GAG and water content
Slow diffusion fraction vs. cellularity
MRI of acellular matrix-based bladder tissue engineering
5
INTRODUCTION
The urinary bladder and bladder disorders
The primary function of the urinary bladder is to store urine at low pressure and
allow voluntary voiding. These functions are regulated by nerves in the spine and around
the bladder and depend on the unique composition and structure of the bladder wall. In a
normal bladder, the wall is composed primarily of three layers: the urothelium, the lamina
propria, and the detrusor muscle. The innermost urothelium consists of highly specialized
cells that provide a barrier to urine penetration into underlying tissues and bloodstream,
while the lamina propria is a connective tissue matrix that limits bladder wall distension
and overall compliance. The detrusor layer consists of smooth muscle cell bundles that are
coordinated by neural control to relax and contract during the filling-voiding cycle, thus
maintaining low pressure and protecting the kidneys from damage.
Bladder dysfunction can result from a variety of congenital and acquired conditions
and compromises quality of life for approximately 400 million people worldwide (1). In
children, the causes are often congenital and include bladder exstrophy (1 in 40,000),
urethral defects (1 in 500) leading to obstructed urine flow, and neuropathic conditions such
as myelomeningocele (1 in 800) that alter neuronal control of the bladder. Loss of proper
bladder control, with subsequent changes in function, can arise from other disruptions to
the neural pathways between the bladder and the micturition center in the brain. In adults,
for example, neurogenic bladders are often induced from trauma to the spinal cord or from
tumors. Obstructive uropathy can also present in adults for a variety of reasons, including
stone formation, enlarged prostates, or tumors. Regardless of the cause of dysfunction, the
MRI of acellular matrix-based bladder tissue engineering
6
structure, thickness, and biomechanics of the bladder wall can be significantly altered. The
result is a high-pressure, low-compliant bladder leading to incontinence and, in severe cases
where pressure extends up the ureters, swelling and impaired function of the kidneys.
The current treatment strategy for bladder dysfunction uses gastrointestinal
segments for replacing bladder tissue (2). This approach can be problematic because
gastrointestinal tissue absorbs solutes that urinary tissue excretes. Due to this difference in
tissue function, several complications can arise, such as infection, metabolic disturbances,
urolithiasis, perforation, and malignancy (2). Alternative methods and materials have been
investigated, most notably tissue expansion (3) and free grafts made from natural (e.g.
small intestinal submucosa, bladder submucosa) or synthetic (e.g. gelatin sponge, polyvinyl
sponge) materials (4,5). However, these attempts have met with limited success due to
biocompatibility, mechanical, structural, and functional problems. Thus, the use of
gastrointestinal segments has remained the gold standard despite its limitations.
Tissue engineering for bladder replacement
Alternative treatments for bladder dysfunction involving tissue engineering
techniques have been pursued in an effort to overcome limitations associated with the
conventional use of gastrointestines. Promising results have been reported in patients using
autologous cells seeded on a collagen-polymer scaffold (6) and in dogs using cell-seeded
allogeneic acellular bladder matrices (7). Prompt angiogenesis of thick tissues is critical to
maintain viability and support continued growth and long-term survival (6,8). Cell-seeding
also is required to ensure tissue regeneration beyond a distance of 0.5 cm (9) necessary for
organ engineering. Compared to the unseeded scaffold, cell seeding reduces fibrosis and
MRI of acellular matrix-based bladder tissue engineering
7
graft contracture and improves overall cellular organization (7). Another requirement is the
use of a scaffold to guide 3D tissue formation. Both synthetic materials and biological
matrices have been used. However, synthetic materials usually succumb to mechanical
failure (6), and biological matrices are believed better able to reproduce the required
biomechanics and also offer intrinsic biochemical characteristics that are difficult to
achieve with synthetic materials. A promising biological scaffold material is the acellular
matrix (ACM) derived from the extracellular matrix of the desired tissue type. The ACM
has cellular components removed to reduce immunogenicity (10) and contains site-
preferred structural and functional molecules for reconstructing tissue. Degradation of the
ACM yields products that have the required angiogenic (11), chemotactic (11,12), and
mitogenic (13) activities to facilitate tissue remodeling. Specifically, these products contain
chemotactic and mitogenic factors that increase recruitment and proliferation of
undifferentiated cells and inhibit the proliferation of differentiated cells. ACMs have been
derived from different tissues, such as skin (14), liver (15), heart valves (16), and bladder
(17), differing in their 3D structure but sharing very similar biochemical composition (13).
Despite early promise, much remains to be accomplished toward engineering a fully
functional bladder. Obstacles include lack of prompt angiogenesis, adverse host-tissue
reaction, inadequate mechanical properties, and lack of innervation. Angiogenesis must be
established promptly for cell expansion and to avoid scarring, but it is currently very
difficult to achieve functional vessels that are long-lasting (18). Another interference with
regeneration is adverse host-tissue reaction, such as that arising from immunogenic
response of cell components or nondegraded scaffold biomaterials. This reaction can be
mitigated with the use of ACM and scaffold manipulation to prevent urine penetration and
MRI of acellular matrix-based bladder tissue engineering
8
its toxic effects (19). Achieving proper biomechanics presents even greater challenges.
Evidence exists that in-vitro mechanical conditioning to simulate the in-vivo environment
can improve tissue morphogenesis, proliferation, differentiation, and function (20,21,22).
However, the type and degree of physical stimuli required and their role in the bladder
remain to be determined. Finally, innervation is an important component to bladder
function but is the least studied and understood. Strategies for nerve regeneration from
other applications perhaps may be applied, such as a recent demonstration of innervating
muscular tissue by seeding primary cultured neurons on a biological extracellular matrix
(23).
Imaging of the tissue engineered bladder
Technology for non-invasive assessment such as MRI is recognized to enable
accelerated progress in tissue engineering and eventual longitudinal monitoring of patients
with tissue engineered organs. Current regeneration approaches need to look beyond
standard histology for a more complete assessment of the function, structure, and
composition of the developing tissue throughout its entire lifespan, from in-vitro cell
seeding to long-term functioning in vivo, in order to identify more effective strategies. In
the bladder, important parameters include cell growth, distribution, viability, and host-
implant interaction in addition to ones more specific to large soft-tissue organs, particularly
those with a mechanical function. These more specific parameters include biomechanics,
functioning of an induced vascular supply, properties of new natural scaffolds for soft-
tissue regeneration, and cell-scaffold interaction.
MRI studies of the tissue engineered bladder have been few but have focused on
MRI of acellular matrix-based bladder tissue engineering
9
several different aspects of regeneration. The earliest efforts addressed the importance of
angiogenesis, investigating the potency of vascular endothelial growth factor (VEGF) and
the ability of MRI to quantify neovessel formation (24,25,26). MRI measures of vascularity
were shown to correlate with increased blood volume at higher VEGF doses (24), and
quantification on an absolute scale was feasible using a blood-pool contrast agent (25).
These studies also demonstrated that enhanced vascularization reduced fibrosis and
improved overall cell repopulation in vivo (26).
The ACM scaffold has been the other main focus because of its role in regeneration,
specifically, in providing an environment conducive to angiogenesis, tissue deposition, and
restoration of structure and function specific to the grafted site (27). MRI characterization
of the ACM has been lacking until recently but is important for establishing a baseline
against which to gauge subsequent tissue development. In a couple of recent MRI studies of
the bladder ACM (28,29), new imaging observations revealed unanticipated biological
phenomena that may be relevant in other ACM-based systems. One important finding was
matrix collagen degradation noted very early on after cell-seeding, possibly due to easier
breakdown of biological tissue compared to synthetic materials (28). Degradation
byproducts are known to modulate tissue remodeling (13), and matrix degradation
concurrent with cell growth is expected in many regeneration paradigms. Distinction of
these concurrent processes using more specific MRI techniques, including multicomponent
T2 as demonstrated in Ref (28), will be critical to accurate assessment. Another valuable
insight is that scaffold optimization from a single manipulation can produce concurrent
biophysical and biochemical changes that have unpredictable effects on MRI parameters.
Specifically, the incorporation of hyaluronic acid, a naturally occurring macromolecule that
MRI of acellular matrix-based bladder tissue engineering
10
can act as a carrier of bioactive molecules and growth factors and has shown potential in
the development of engineered tissues (30,31), was shown to increase macromolecular
content and ACM hydration (29). These concurrent changes exerted competing effects on
MRI tissue relaxation and diffusion, and single MRI parameters failed to provide adequate
specificity. These studies (28,29) highlight the ability of MRI to improve our understanding
of uncharted development of engineered tissues by detecting changes for further
investigation and validation. They also emphasize the need for more specific MRI methods
to assess tissue structure, composition, and function.
In this study, we explore MRI methods for improved specificity in monitoring tissue
development in a bladder ACM-based regeneration model. Multiexponential diffusion and
effective transverse relaxation time, T2*, are investigated for their ability to discriminate
cellularization and matrix changes without influence from concurrent or undesired sources.
EXPERIMENTAL
Acellular matrix (ACM) preparation
Acellular tissue matrices were prepared according to a published protocol (10).
Fresh urinary bladders were harvested from pigs (20-50 kg) and immediately washed in
sterile phosphate buffer saline (Sigma), longitudinally sectioned, and placed in a hypotonic
solution [5 mM EDTA , 10 mM Tris HCl, pH 8.0, 1% Triton X-100, 0.1 mg/mL Pefabloc
PlusTM
(Alexis) and Penicillin/Streptomycin] (Sigma) for 72 hours, stirring at 4˚C, to lyse
all cell structures and inhibit proteases. Bladder tissues were then immersed in hypertonic
solution [5 mM EDTA, 10 mM Tris HCl, pH 8.0, 1% Triton X-100 and 1.5 M KCl]
MRI of acellular matrix-based bladder tissue engineering
11
(Sigma), stirring for an additional 72 hours at 4˚C to denature residual proteins.
Subsequently, bladders were washed in Hank’s Balanced Salt Solution (HBBS), followed
by overnight incubation at 37˚C with Benzonase (2U/mL) diluted in HBSS to degrade
DNA and RNA components. Final extraction was performed by transferring bladders to a
0.25% CHAPS (non-denaturing detergent) based solution [50 mM Tris HCl, pH 8.0, 1%
Triton X-100 and Penicillin/Streptomycin]. The resulting ACM was washed with sterile
double distilled water (dH2O) and stored in 70% ethanol. Hematoxylin and eosin (H&E)
sections were taken to confirm acellularity. All protocols were approved by the institutional
Animal Care Committee and compliant with national policies on the humane use of
laboratory animals (CCAC guidelines).
Preparation of ACM and cell-seeded constructs
ACMs of 3 mm thickness were cut into 22 cm2 squares, dehydrated by gradual
immersion in increasing concentrations of ethanol (50%, 70%, 80%, 90%, 100%) for 1
hour each at room temperature, and lyophilized for 24 hours (VirTis-temp, -70˚C and
vacuum, 120 millitorr). Hyaluronic acid (HA) was incorporated by rehydrating ACMs in
increasing concentrations of HA (0.05, 0.1, 0.2 mg/100 mL) (Sigma) shaking for one hour,
then in 0.5 mg/100 mL HA for 24 hours, all at 37˚C. Excess HA was washed off with
dH2O. Alcian blue staining was performed to confirm HA uptake (Figure 1). ACMs with
and without HA were immersed in medium (high glucose Dulbecco’s modified essential
medium (Wisent Inc), 10% fetal bovine serum) at 4°C.
For cell-seeding, cell cultures were obtained from passage 2 smooth muscle cells
isolated from porcine bladder (32) and expanded in medium. Cells were seeded on ACMs
MRI of acellular matrix-based bladder tissue engineering
12
in a 6 well plate at 1106
cells/cm2
in 100 L of medium. The cells were allowed to attach
on the ACM for 3 hrs (37°C in a humidified atmosphere of 5% CO2) before incubating in 2
mL of medium. The medium was changed every 72 hours.
Experimental design
Cell-seeded (Ns=20) and unseeded (Nu=15) ACMs were prepared as described
above. Six of the unseeded ACMs were further processed for HA incorporation. Imaging
was performed on day 1, 3, and 7 and, for seeded ACMs, at an additional timepoint of 21
days after cell-seeding. Samples were prepared for MRI by transferring each ACM to a 5
mL, 12 mm diameter round Falcon tube containing medium, and placing the tubes in a
water-filled container. Histology was also performed, and samples were taken prior to
imaging to verify no changes occurred during MRI. Additional validation measurements
involving immunofluorescence microscropy and biochemical assays from previous
experiments are reported here to provide a benchmark.
Histology
All tissue samples were fixed in 10% formalin for 24 hours, embedded in paraffin,
and sectioned into 5 m thick slices. Sections were stained with H&E and Masson’s
Trichrome and examined under a light microscope (Nikon Eclipse E 400). Cell density was
assessed qualitatively at all time-points on H&E sections. Four different classes of
cellularity were defined: I (very few cells), II (few cells), III (patches of cells), and IV
(good cellularity). A blinded histology assessment was performed by a pathologist.
MRI of acellular matrix-based bladder tissue engineering
13
Quantitative MRI
MRI was performed on a 1.5 Tesla clinical scanner (Signa EXCITE TwinSpeed, GE
Healthcare, Milwaukee, WI) using a single 3-inch surface coil positioned under a water-
filled container in which ACM-containing tubes were placed. Pilot scans were taken to
determine placement of imaging slices perpendicular to the tubes and encompassing the
upper and lower ends of the ACMs.
Quantitative T1, T2, T2*, and diffusion imaging was performed. The longitudinal
relaxation time T1 was measured using a rapid 3D technique (33) based on a fast spoiled
gradient echo sequence with radio-frequency field correction [flip angle = 2, 3, 10, 20,
number of averages (NAVG) = 4]. The transverse relaxation time T2 was measured using a
2D spin echo sequence [9 echoes with echo times (TE) = 9 - 300 ms, repetition time (TR) =
3000 ms, NAVG = 1]. The effective transverse relaxation time T2* was measured using a 2D
multi-echo gradient-echo sequence [16 equally spaced TEs = [2.6 – 34.4] ms, TR = 118 ms,
FA = 20, NAVG = 4]. Diffusion imaging was performed using a spin echo diffusion-
weighted imaging sequence up to high b-values [b = 0 – 3000 s/mm2
in steps of 200 s/mm2,
TR = 4000 ms, NAVG = 16]. In addition to high b-value acquisition for multiexponential
diffusion analysis, single diffusion coefficients were also obtained by imaging at a b-value
of 1000 s/mm2. Slice thickness was 3 mm for all sequences; in-plane resolutions were 0.4
mm (T1, T2, T2*) and 0.8 mm for diffusion.
All image post-processing of quantitative T1, T2, and T2* data was performed on a
pixel-wise basis using in-house scripts developed in Matlab (V.7.0, Mathworks Inc.,
Natick, MA, USA). T1 parameter maps were generated as previously described, with
MRI of acellular matrix-based bladder tissue engineering
14
analytical-based flip angle correction using B1 field maps acquired separately (33). Both
T2 and T2* parameter maps were computed by fitting a mono-exponential signal decay
function to signal intensities obtained at various echo times (34). The mean T1, T2, or T2*
value for each ACM was then obtained by averaging within the ACM and through all
imaging slices through which the ACM traversed. For diffusion data, a region
encompassing the ACM was drawn and the average signal intensity at each b-value was
taken for analysis. Non-linear least squares analysis, which requires no a priori knowledge
of the number of the exponentials, was used to estimate diffusion coefficients and their
fractions. All data are expressed as mean value ± standard deviation.
Statistical analysis
Differences between groups were assessed using a two-tailed Student’s t-test.
Significance was declared at P < 0.05.
RESULTS
Histology
Cellularity varied over the time interval of 1 to 21 days post-cell seeding, with cells
aggregated mainly at the surface of the ACM. Representative H&E sections showing the
degree of cellularity associated with each of the four classifications are given in Figure 2.
No difference in cellularity was observed in samples taken before and after MRI from the
same ACM, indicating that the time spent for MRI did not affect cell viability.
Masson’s Trichome sections (Figure 3) illustrate the main constituents of the ACM:
smooth muscle bundles within a collagen extracellular matrix. Histologically, the ACM
MRI of acellular matrix-based bladder tissue engineering
15
appears consistent amongst the different preparations: cell-seeding and HA incorporation.
However, it is known from previous immunohistochemistry and biochemistry experiments
that cell-seeding results in matrix collagen degradation from cell-released collagenase (28),
and HA incorporation increases both macromolecular content and tissue hydration (29).
Table 1 summarizes these biochemical changes. These concurrent changes present
conflicting influences on single MRI parameters such as T1 and T2 and thereby limit their
specificity. For example, increased T1 and, to a much greater extent, increased T2 from
hydration can confound our assessment of cell growth or matrix glycosaminoglycan (GAG)
content. The T2 spectrum was previously shown to distinguish these concurrent changes
(Figure 4), but its assessment of biophysical boundaries (such as cells) would be limited
due to its sensitivity to biochemical composition.
Quantitative MRI
Examples MR images of the construct are shown in Figure 5. From high b-value
diffusion imaging, sensitivity to presence of cells in the ACM is shown. Figure 6 shows
that cell-seeded and unseeded ACMs exhibit biexponential and monoexponential diffusion
decay, respectively. This behaviour is consistent with a fast diffusion component associated
with the extracellular compartment and a slow diffusion component associated with the
intracellular space. Figure 7 shows that the slow diffusion fraction has the highest
correlation with cellularity (r = 0.954, P < 10-5
) compared to single diffusion coefficient (r
= -0.153, P = 0.654), T1 (r = -0.614, P < 0.05), and T2 (r = -0.309, P = 0.355).
New observations on T2*, which is predominantly used in tissue iron and oxygen
measurements, suggest its potential role as a more specific parameter for assessment of
MRI of acellular matrix-based bladder tissue engineering
16
tissue composition. Figure 8 shows that cell-seeding produces the greatest relative change
on T2* (P < 10-4
) compared to T2 (P < 0.05) or T1 (P = 0.11), although the correlation
between T2* and cellularity was not significant (r = 0.152, P = 0.676). Figure 9 shows that
the presence of the macromolecule HA in the unseeded ACM produces significant changes
on T2*, T2, and T1 (P < 0.05), but the largest relative change is seen on T2* and appears to
be due to reduced sensitivity to tissue hydration. Note that while both T1 and T2 changed in
the same direction with cell-seeding or incorporation of HA, only T2* changed in opposing
directions with these two manipulations.
DISCUSSION
Improved specificity in MRI measurements was investigated in this study for
monitoring tissue development in a bladder ACM-based paradigm for soft-tissue
regeneration. The ACM as a biological scaffold is a relatively new approach for guiding 3D
tissue formation and is especially promising for whole organ tissue engineering, especially
those with a mechanical function. Aside from reducing an immunogenic response, the
ACM possesses the required structural and mechanical properties and its degradation
releases by-products that facilitate tissue remodeling. This approach may provide truly off-
the-shelf replacement tissue if animal-derived ACMs can be used in humans. For instance,
given the similar composition and biomechanical properties between porcine and human
ACMs (35), some investigators have taken the next step and seeded human bladder cells on
porcine ACM (36). Nonetheless, experience with biological matrices is in early stages, and
there are few related imaging studies. The first MRI studies of ACM-based soft-tissue
regeneration reported early matrix degradation (28), which is not seen or probably arises
MRI of acellular matrix-based bladder tissue engineering
17
much later in conventional scaffold materials. This concurrency of matrix degradation and
cell growth can confound assessment of either process if common MRI parameters such as
T1, T2, and single diffusion coefficient are used. Also, tissue hydration can vary
substantially with matrix manipulation (29), likely much more so than in synthetic polymer
scaffolds. Because of these properties, the ACM may require more specific MRI techniques
to distinguish relevant processes amidst less important concurrent changes, such as those
related to tissue water content.
For the assessment for cell growth, this study shows that multiexponential diffusion
analysis may provide the best specificity by minimizing sensitivity to other changes, such
as matrix degradation or tissue hydration. These other changes exert an appreciable effect
on more commonly used parameters in tissue engineered systems, such as T1, T2, and
single diffusion coefficient (37), and underlie the weak correlation between these
parameters and cellularity as observed in this study. Amongst these three parameters, only
T1 exhibits a trend (decrease) consistent with cell growth. The other two parameters, T2
and diffusion, are more unpredictable probably due to their greater sensitivity to water
content and changes in the extracellular matrix structure. In contrast, the slow diffusion
component in multiexponential diffusion decay is sensitive to the intracellular compartment
and quite insensitive to changes in the extracellular environment. It also provides a more
specific technique to measure cellularity without confounders from tissue biochemistry.
New properties on the effective transverse relaxation time T2* were observed in this
study and suggest improved specificity over T2 measurements. One consistent observation
was that unlike T2, T2* was relatively insensitive to water content. This is best appreciated
by comparing ACMs with and without HA (Figure 9). In principle, macromolecules
MRI of acellular matrix-based bladder tissue engineering
18
including proteins induce a lowering in T2 and, therefore, T2* compared to free water. This
is due to more restricted water in the hydration layer on the surface of the macromolecules.
However, being hydrophilic in nature, HA also elevates tissue hydration and, thus, free
water content, therefore increasing T2. The competing T2 effects from a higher GAG
content and increased hydration resulted in a net increase, whereas T2* remained low. The
only plausible explanation is that T2* is much less sensitive to water content, likely
because an increase in free water does not introduce local centers of magnetic non-
uniformities; thus, T2* dephasing is not affected. This reduced sensitivity of T2* to
hydration was speculated in a recent study of chondrocyte transplantation in patients (38),
but histological validation was lacking. In this study, biochemistry validation confirms that
T2* is much more specific to macromolecular content than T2 or T1. A second observation
in this study was a substantially larger T2* in cell-seeded relative to unseeded ACMs
(Figure 8). This result is more difficult to interpret. A possible explanation is that cell-
seeding, which is known to fill pores of the ACM, reduces T2* dephasing associated with
pore interfaces. This result is particularly relevant in tissue engineering applications,
because scaffolds are designed to be porous for the purpose of embedding cells. The ability
to identify when cells have occupied these pores is very valuable for determining how
firmly entrenched cells are.
While this study has suggested new potential in tissue engineering applications for
multiexponential diffusion and T2* measurements, our results are consistent with MRI
studies in other applications that attempt to elucidate underlying biological mechanisms.
For example, diffusion studies of perfused breast cancer cells in vitro showed that a second
slow diffusion component associated with restricted intracellular diffusion arose when cells
MRI of acellular matrix-based bladder tissue engineering
19
were added to a gel medium (39). Other studies have shown a correlation between the slow
diffusion fraction and intracellular space in cell suspensions (40,41). However, caution
must be taken in interpreting the slow diffusion fraction in terms of absolute cellular
volume. The biexponential diffusion model is valid only when there is no exchange
between intra- and extracellular water compartments and no restriction of intracellular
water by cell membranes. When these conditions are unmet, diffusion decay deviates from
a pure biexponential, and the diffusion fractions do not correspond to the actual sizes of the
two pools (42,43,44). In order to achieve the ideal situation, water exchange must be very
slow, such as when there are short diffusion times, slower diffusion rates, large cells, or low
membrane permeability. Nonetheless, the slow diffusion fraction provides a useful metric
to estimate cellularity even if not on an absolute scale.
Our T2* findings are more challenging to interpret, as there are relatively few
literature comparisons. Nonetheless, our hypothesis that increased T2* is due to cell filling
of matrix pores, decreased T2* to higher macromolecular content, and relative insensitivity
of T2* to hydration are plausible given the known behaviour of T2* decay. The association
with pores, however, requires more explanation. Materials with pores larger than 15Ǻ
exhibit increased transverse relaxation due to the ability of large pores to take up water
(45). Scaffolds contain much larger pores, in the microns range, that are expected to contain
free water and give rise to susceptibility differences at pore interfaces. This difference leads
to T2* dephasing, the magnitude of which is likely lessened when pores are replaced with
cells.
There are a number of different avenues for future investigation, and new
challenges will present themselves as the regeneration strategy becomes increasingly
MRI of acellular matrix-based bladder tissue engineering
20
complex. For instance, matrix- and cell-related changes, which can exert opposing or
constructive influences, may need to be distinguished. Also, other essential layers of the
bladder such as the urothelium and lamina propria need to be formed, and it remains to be
determined if cells for these layers can be recruited in vivo and not require direct seeding as
does the smooth muscle layer. To identify these multiple signatures will be a challenge,
likely to require finding matrix- or cell-type-specific molecular signatures that can be
detected on MRI. Improvement can also be made in assessing cell repopulation of the
construct. First, the multiexponential diffusion method for measuring cellularity would
benefit from imaging at higher field strengths, which allows shorter diffusion times and
approach to the slow exchange limit. Measuring the number of viable cells would be even
more informative, and this may be achieved through 1H spectroscopy, where a linear
correlation between cell number and the choline peak has been previously observed
(46,47). Quantitative validation of cell number counts (e.g. DNA quantification) will be
undertaken in future studies. Further investigation is also needed to understand the T2*
changes observed in this study. Our hypothesis that T2* may identify cell filling of pore
space should be evaluated in polymer scaffolds where pore size and number can be
systematically controlled. Another finding related to T2*, namely, its insensitivity to tissue
hydration that was also observed in chondrocyte transplantation (38), is potentially valuable
in a wide variety of applications. For example, inflammation, which often results from
tissue construct-host interaction, should be investigated in the future to determine if T2* is
sensitive to infiltration of inflammatory bodies but not to accompanying edema.
In conclusion, this study has demonstrated the potential role of multiexponential
diffusion and T2* for more specific assessment of cell growth and tissue development. The
MRI of acellular matrix-based bladder tissue engineering
21
results of this study are relevant for regeneration paradigms based on biological scaffolds
such as the acellular matrix for engineering soft tissue and large organs. They also extend to
other applications where concurrent biological changes exist, for example, where cellularity
needs to determined independent of biochemical changes or where tissue hydration is a
confounder and needs to be removed.
MRI of acellular matrix-based bladder tissue engineering
22
REFERENCES
1 Korossis S, Bolland F, Ingham E, Fisher J, Kearney J, Southgate J. Review: tissue
engineering of the urinary bladder: considering structure-function relationships and
the role of mechanotransduction. Tissue Eng. 2006; 12: 635-644.
2 Atala A. Bioengineered Tissues for urogenital repair in children. Ped. Res. 2008; 63:
569-575.
3 Satar N, Yoo JJ, Atala A. Progressive dilation for bladder tissue expansion. J. Urol.
1999; 162: 829-831.
4 Vaught JD, Kropp BP, Sawyer BD, Rippy MK, Badylak SF, Shannon HE, Thor KB.
Detrusor regeneration in the rat using porcine small intestinal submucosal grafts:
functional innervation and receptor expression. J. Urol. 1996; 155: 374-378.
5 Kudish HG. The use of polyvinyl sponge for experimental cystoplasty. J. Urol. 1957;
78: 232-235.
6 Atala A, Bauer SB, Soker S, Yoo JJ, Retik AB. Tissue-engineered autologous
bladders for patients needing cystoplasty. Lancet 2006; 367: 1241-1246.
7 Oberpenning F, Meng J, Yoo JJ, Atala A. De novo reconstitution of a functional
mammalian urinary bladder by tissue engineering. Nat. Biotechnol. 1999; 17: 149-
155.
8 Mooney DJ, Mikos AG. Growing new organs. Sci. Am. 1999; 280: 60-65.
9 Dorin RP, Pohl HG, De Filippo RE, Yoo JJ, Atala A. Tubularized urethral
replacement with unseeded matrices: what is the maximum distance for normal tissue
regeneration? World J. Urol. 2008; 26: 323-326.
MRI of acellular matrix-based bladder tissue engineering
23
10 Brown AL, Farhat W, Merguerian PA, Wilson GJ, Khoury AE, Woodhouse KA. 22
week assessment of bladder acellular matrix as a bladder augmentation material in a
porcine model. Biomaterials 2002; 23: 2179-2190.
11 Li F, Li W, Johnson S, Ingram D, Yoder M, Badylak S. Low-molecular-weight
peptides derived from extracellular matrix as chemoattractants for primary endothelial
cells. Endothelium 2004; 11: 199-206.
12 Zantop T, Gilbert TW, Yoder MC, Badylak SF. Extracellular matrix scaffolds attract
bone marrow derived cells in a mouse model of Achilles tendon reconstruction. J.
Orthop. Res. 2006; 24: 1299-1309.
13 Reing JE, Zhang L, Myers-Irvin J, Cordero KE, Freytes DO, Heber-Katz E,
Bedelbaeva K, McIntosh D, Dewilde A, Braunhut SJ, Badylak SF. Degradation
products of extracellular matrix affect cell migration and proliferation. Tissue Eng.
Part A 2009; 15: 605-614.
14 Armour AD, Fish JS, Woodhouse KA, Semple JL. A comparison of human and
porcine acellularized dermis: interactions with human fibroblasts in vitro. Plast.
Reconstr. Surg. 2006; 117: 845-856.
15 Brown B, Lindberg K, Reing J, Stolz DB, Badylak SF. The basement membrane
component of biologic scaffolds derived from extracellular matrix. Tissue Eng. 2006;
12: 519-526.
16 Lichtenberg A, Tudorache I, Cebotari S, Suprunov M, Tudorache G, Goerler H, Park
JK, Hilfiker-Kleiner D, Ringes-Lichtenberg S, Karck M, Brandes G, Hilfiker A,
MRI of acellular matrix-based bladder tissue engineering
24
Haverich A. Preclinical testing of tissue-engineered heart valves re-endothelialized
under simulated physiological conditions. Circulation 2006; 114: I559-I565.
17 Chen F, Yoo JJ, Atala A. Acellular collagen matrix as a possible ‘‘off the shelf ’’
biomaterial for urethral repair. Urology 1999; 54: 407-410.
18 Jain RK. Molecular regulation of vessel maturation. Nat. Med. 2003; 9: 685-693.
19 Cartwright LM, Shou Z, Yeger H, Farhat WA. Porcine bladder acellular matrix
porosity: impact of hyaluronic acid and lyophilization. J. Biomed. Mater. Res. A
2006; 77: 180-184.
20 Ingber DE, Dike L, Hansen L, Karp S, Liley H, Maniotis A, McNamee H, Mooney D,
Plopper G, Sims J, et al. Cellular tensegrity: exploring how mechanical changes in the
cytoskeleton regulate cell growth, migration, and tissue pattern during
morphogenesis. Int. Rev. Cytol. 1994; 150: 173-224.
21 Oluwole BO, Du W, Mills I, Sumpio BE. Gene regulation by mechanical forces.
Endothelium 1997; 5: 85-93.
22 Grinnell F. Fibroblasts, myofibroblasts, and wound contraction. J. Cell Biol. 1994;
124: 401-404.
23 Agrawal V, Brown BN, Beattie AJ, Gilbert TW, Badylak SF. Evidence of innervation
following extracellular matrix scaffold-mediated remodelling of muscular tissues. J.
Tissue Eng. Regen. Med. 2009; 3: 590-600.
MRI of acellular matrix-based bladder tissue engineering
25
24 Cheng HL, Chen J, Babyn PS, Farhat WA. Dynamic Gd-DTPA enhanced MRI as a
surrogate marker of angiogenesis in tissue-engineered bladder constructs: a feasibility
study in rabbits. J. Magn. Reson. Imaging 2005; 21: 415-423.
25 Cheng HL, Wallis C, Shou Z, Farhat WA. Quantifying angiogenesis in VEGF-
enhanced tissue-engineered bladder constructs by dynamic contrast-enhanced MRI
using contrast agents of different molecular weights. J. Magn. Reson. Imaging 2007;
25: 137-145.
26 Cartwright L, Farhat WA, Sherman C, Chen J, Babyn P, Yeger H, Cheng HL.
Dynamic contrast-enhanced MRI to quantify VEGF-enhanced tissue-engineered
bladder graft neovascularization: pilot study. J. Biomed. Mater. Res. A 2006; 77: 390-
395.
27 Hodde J. Naturally occurring scaffolds for soft tissue repair and regeneration. Tissue
Eng. 2002; 8: 295-308.
28 Cheng HL, Islam SS, Loai Y, Antoon R, Beaumont M, Farhat WA. Quantitative MRI
assessment of matrix development in cell-seeded natural urinary bladder smooth
muscle tissue-engineered constructs. Tissue Eng. Part C Methods 2010 Apr 13 [Epub
ahead of print]
29 Cheng HL, Islam SS, Loai Y, Antoon R, Beaumont M, Farhat WA. The acellular
matrix (ACM) for bladder tissue-engineering: a quantitative magnetic resonance
imaging study. Magn. Reson. Med. 2010 May 25 [Epub ahead of print]
MRI of acellular matrix-based bladder tissue engineering
26
30 Chung C, Erickson IE, Mauck RL, Burdick JA. Differential behavior of auricular and
articular chondrocytes in hyaluronic acid hydrogels. Tissue Eng. Part A 2008;
14:.1121-1131.
31 Zavan B, Vindigni V, Lepidi S, Iacopetti I, Avruscio G, Abatangelo G, Cortivo R.
Neoarteries grown in vivo using a tissue-engineered hyaluronan-based scaffold.
FASEB J. 2008; 22: 2853-2861.
32 Farhat WA, Chen J, Sherman C, Cartwright L, Bahoric A, Yeger H. Impact of fibrin
glue and urinary bladder cell spraying on the in-vivo acellular matrix cellularization: a
porcine pilot study. Can. J. Urol. 2006; 13: 3000-3008.
33 Cheng HL, Wright GA. Rapid high-resolution T(1) mapping by variable flip angles:
accurate and precise measurements in the presence of radiofrequency field
inhomogeneity. Magn. Reson. Med. 2006; 55: 566-574.
34 Beaumont M, Odame I, Babyn PS, Vidarsson L, Kirby-Allen M, Cheng HL. Accurate
liver T2 measurement of iron overload: a simulations investigation and in vivo study.
J. Magn. Reson. Imaging 2009; 30: 313-320.
35 Dahms SE, Piechota HJ, Dahiya R, Lue TF, Tanagho EA. Composition and
biomechanical properties of the bladder acellular matrix graft: comparative analysis
in rat, pig and human. Br. J. Urol. 1998; 82: 411-419.
36 Ram-Liebig G, Ravens U, Balana B, Haase M, Baretton G, Wirth MP. New
approaches in the modulation of bladder smooth muscle cells on viable detrusor
constructs. World J. Urol. 2006; 24: 429-437.
MRI of acellular matrix-based bladder tissue engineering
27
37 Peptan IA, Hong L, Xu H, Magin RL. MR assessment of osteogenic differentiation in
tissue-engineered constructs. Tissue Eng. 2006; 12: 843-851.
38 Welsch GH, Trattnig S, Hughes T, Quirbach S, Olk A, Blanke M, Marlovits S,
Mamisch TC. T2 and T2* mapping in patients after matrix-associated autologous
chondrocyte transplantation: initial results on clinical use with 3.0-tesla MRI. Eur.
Radiol. 2010; 20: 1515-1523.
39 Van Zijl PC, Moonen CT, Faustino P, Pekar J, Kaplan O, Cohen JS. Complete
separation of intracellular and extracellular information in NMR spectra of perfused
cells by diffusion-weighted spectroscopy. Proc. Natl. Acad. Sci. U S A 1991; 88:
3228-3232.
40 Thelwall PE, Grant SC, Stanisz GJ, Blackband SJ. Human erythrocyte ghosts:
Exploring the origins of multiexponential water diffusion in a model biological tissue
with magnetic resonance. Magn. Reson. Med. 2002; 48: 649-657.
41 Roth Y, Ocherashvilli A, Daniels D, Ruiz-Cabello J, Maier SE, Orenstein A, Mardor
Y. Quantification of water compartmentation in cell suspensions by diffusion-
weighted and T(2)-weighted MRI. Magn. Reson. Imaging 2008; 26: 88-102.
42 Frøhlich AF, Ostergaard L, Kiselev VG. Effect of impermeable boundaries on
diffusion-attenuated MR signal. J. Magn. Reson. 2006; 179: 223-233.
43 Li JG, Stanisz GJ, Henkelman RM. Integrated analysis of diffusion and relaxation of
water in blood. Magn. Reson. Med. 1998; 40: 79-88.
MRI of acellular matrix-based bladder tissue engineering
28
44 Chin CL, Wehrli FW, Hwang SN, Takahashi M, Hackney DB. Biexponential
diffusion attenuation in the rat spinal cord: Computer simulations based on anatomic
images of axonal architecture. Magn. Reson. Med. 2002; 47: 455-460.
45 Watanabe T, Murase N, Staemmler M, Gersonde K. Multiexponential proton
relaxation processes of compartmentalized water in gels. Magn. Reson. Med. 1992;
27: 118-134.
46 Stabler CL, Long RC, Sambanis A, Constantinidis I. Noninvasive measurement of
viable cell number in tissue-engineered constructs in vitro, using 1H nuclear magnetic
resonance spectroscopy. Tissue Eng. 2005; 11: 404-414.
47 Stabler CL, Long RC Jr, Constantinidis I, Sambanis A. In vivo noninvasive
monitoring of a tissue engineered construct using 1H NMR spectroscopy. Cell
Transplant 2005; 14: 139-149.
MRI of acellular matrix-based bladder tissue engineering
29
Table 1
Biophysical and biochemical changes in ACM from different manipulations
Manipulation Tissue Changes
Incorporation of
hyaluronic acid (HA)
Total GAG concentration increases, with a 2:1 ratio of non-
sulphated (i.e. HA) versus sulphated GAG.
Water content nearly doubles.
Seeding with smooth
muscle cells
Matrix collagen I degrades
MRI of acellular matrix-based bladder tissue engineering
30
FIGURE LEGENDS
Figure 1. Alcian blue staining confirms the uptake of hyaluronic acid (HA) in HA-ACM
constructs, which have a blue hue compared with ACM constructs without HA. (magnification,
10).
Figure 2. Representative H&E stained sections for the different classifications of cellularity: I
(very few cells), II (few cells), III (patches of cells), and IV (good cellularity). (magnification,
10).
Figure 3. Masson’s Trichome staining of the ACM shows a collagen extracellular matrix and
smooth muscle bundles (upper left corner). (magnification, 10).
Figure 4. Changes in the T2 spectrum (solid to dotted curves) can distinguish a) increased tissue
water content accompanying incorporation of hyaluronic acid and b) degradation of the
extracellular matrix with cell-seeding. In a) the higher water content is evident in a shift of the
long T2 component towards higher values. In b) progressive loss of matrix collagen is evident in
a reduced fraction of the short T2 component.
Figure 5. a) Diffusion-weighted and b) T2*-weighted MR images of an ACM in a 12-mm
diameter tube. Diffusion-weighted images are illustrated for b-values of 200 s/mm2 (left) and
1600 s/mm2 (right). T2*-weighted images are illustrated for echo times of 2.6 ms (left) and 34.4
ms (right).
Figure 6. Diffusion decay characteristics are different between unseeded ACMs (filled circles)
and seeded ACMs (open circles). Unseeded ACMs exhibit monoexponential decay, whereas
biexponential behaviour emerges in seeded ACMs due to the presence of both an extra- and
intracellular compartment.
MRI of acellular matrix-based bladder tissue engineering
31
Figure 7. Quantitative MRI parameters for discriminating various levels of cellularity. The slow
diffusion fraction, which represents the proportion of intracellular space, is significantly
correlated with increasing cellularity (I, II, III, IV). The other parameters (single diffusion
coefficient D, T1, and T2) show much lower correlation or none at all.
Figure 8. Effect on MRI parameters from seeding cells on ACM. T2* exhibits a significant and
greater relative change compared to either T2 or T1. The T2* increase is postulated to arise from
cell-filling of matrix pores. Mean values and standard deviation are shown in units of ms for
unseeded (shaded) and seeded (non-shaded) ACMs. ** P < 0.01, * P < 0.05.
Figure 9. Effect on MRI parameters from incorporating hyaluronic acid in ACM. T2* exhibits a
significant and greater relative change compared to either T2 or T1. The T2* decrease is
consistent with higher macromolecular content in the ACM and suggests that unlike T2, T2* is
relatively insensitive to tissue hydration. Mean values and standard deviations are shown in units
of ms for ACM (shaded) and HA-ACM (non-shaded). * P < 0.05.
FIG. 1
HA-ACMACM
FIG. 1
I II
III IV
FIG. 2
FIG. 3
T2 (ms)
0.1
0
0.1
0100 101 102 103
ampl
itude
ampl
itude
a
b
FIG. 5
a
b
0
500
1000
1500
2000
2500
3000
0
50
100
150
200
250
300
280
300
320
340
360
380
400
420
440
280
300
320
340
360
380
400
420
440
FIG. 4
0.01
0.1
1
0 500 1000 1500 2000 2500 3000
b-value (s/mm2)
Rel
ativ
e si
gnal
(S/S
o)
3 300
2
0.2
D (1
0-3
mm
2 /s)
T1 (s
)
T2 (m
s)
1
100
0
2002
1
0
0
0.1
I II III IV I II III IV
Cellularity Cellularity
FIG. 5
Slow
diff
usio
n fr
actio
n
42.7
71.3
125
159 19921854
T2* T2 T1
a.u.
FIG. 8
** *
42.7
23.4
125
156 1992
1526
T2* T2 T1
a.u.
FIG. 9
** *
Recommended