Note: Descriptions are shown in the official language in which they were submitted.
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MICROFLUIDIC DEVICE FOR THE DIGESTION OF TISSUES INTO
CELLULAR SUSPENSIONS
Related Applications
[0001] This Application claims priority to U.S. Patent Application No.
17/180,711, filed
on February 19, 2021, which is a continuation-in-part of U.S. Patent
Application No.
16/115,434, filed on August 28, 2018, now issued as U.S. Patent No.
10,926,257, which also
claims priority to U.S. Provisional Patent Application No. 62/551,172 filed on
August 28,
2017, which are hereby incorporated by reference in their entirety. Priority
is claimed
pursuant to 35 U.S.C. 119, 120 and any other applicable statute.
Statement Re2ardin2 Federally Sponsored
Research and Development
[0002] This invention was made with Government support under Grant No. IIP-
1362165
awarded by the National Science Foundation (NSF). The Government has certain
rights in the
invention.
Technical Field
[0003] The technical field generally relates to microfluidic devices that
are used to digest
tissue specimens or tissue samples into cellular suspensions.
Back2round
[0004] The past decade has seen a rapid growth in interest to harvest
single cells from
tissues that has spanned across several biomedical research areas. This has
been driven in part
by the rise in use of single cell analysis techniques, such as flow cytometry,
mass
spectroscopy, and single cell sequencing, to identify and profile the diverse
cell types
typically found within tissues. For cancer, this has enabled assessment of
tumor
heterogeneity, metastatic potential, and the presence of rare cell types such
as putative cancer
stem cells. These insights obtained at the resolution of single cells are
drastically changing
the understanding of cancer, and in the future are poised to revolutionize
clinical diagnostics
and inform personalized patient care. In the field of tissue engineering,
isolation of primary
cells from tissues is critical for the creation of new constructs to replace
damaged organs,
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such as skin, liver, heart, pancreas and kidney. Finally, a major goal of
regenerative medicine
is to isolate mesenchymal stem cells and progenitor cells from tissues to heal
or otherwise
replace diseased parts of the body. A common theme unifying all of these
applications is that
they require viable single cells that remain as representative of their
original phenotypic state
as possible. Thus, there is a critical need to develop new technologies that
will make it
possible to liberate single cells from tissues in a rapid, gentle, and
thorough manner.
[0005] Microfluidic technologies have emerged as simple yet powerful
methods for
processing and manipulating cellular samples at the microscale. However, only
a few
microfluidic devices have been developed to work with cell aggregates and
tissues. The
microfluidic cell dissociation chip (p.-CDC) described by Lin et al. was
designed to break
down neurospheres under fluid flow using a micro-pillar array. See Lin et al.,
Separation of
Heterogenous Neural Cells in Neurospheres using Microfluidic Chip, Anal Chem,
85, 11920-
8 (2013). However, this device could only be used with aggregates that were
less than 300
p.m in diameter, and yet still suffered from clogging issues. Wallman et al.
disclosed a
Biogrid device that was designed to mechanically cut neurospheres using sharp
silicon knife-
edges placed across the device cross-section. See Wallman et al., Biogrid ¨ a
microfluidic
device for large-scale enzyme-free dissociation of stem cell aggregates, Lab
Chip, 11(19), pp.
3241-8 (2011). While more effective, mechanical cutting in this fashion was
harsh and only
resulted in smaller aggregates, not single cells. In previous work, a
microfluidic device was
disclosed that employed a network of branching channels to achieve highly
efficient and
rapid dissociation of cancer cell aggregates into viable single cells. See Qui
et al.,
Microfluidic device for mechanical dissociation of cancer cell aggregates into
single cells,
Lab on a Chip, 15.1, 339-350 (2015). However, the inlet could not accommodate
samples
that were greater than 1 mm in size, requiring off-chip mincing and digestion
of larger tissue
specimens. While full scale tissues have been employed in a single
microfluidic application,
namely, the culture and enzymatic digestion of rat liver biopsies, this device
has a number of
limitations. See Hattersley et al., Development of a microfluidic device for
the maintenance
and interrogation of viable tissue biopsies, Lab Chip, 8(11), pp. 1842-6
(2008). For example,
this device just provided a means to incubate tissues with enzymes, and
suffered from
extremely low cell yields, even after prolonged digestion times.
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Summary
[0006] In one embodiment, a microfluidic device for the processing of a
tissue sample into
cellular suspensions includes a substrate or chip having formed therein an
inlet, an outlet, and
a sample chamber dimensioned to hold the tissue sample. The sample chamber is
fluidically
coupled at one side to a plurality of upstream hydro-mincing microfluidic
channels disposed
in the substrate or chip. These upstream hydro-mincing microfluidic channels
drive the fluid
into discrete locations of the tissue in a jetting process, effectively
mincing it through the
application of hydrodynamic shear forces and improved enzyme penetration
(contained in the
fluid). The sample chamber is further fluidically coupled at another side of
the sample
chamber to a plurality of downstream sieve microfluidic channels disposed in
the substrate or
chip and further fluidically coupled to the outlet. The downstream sieve
microfluidic
channels act as a sieve that firmly holds the tissue in place while also
allowing smaller
aggregates and cells to exit the sample chamber.
[0007] In some embodiments, the microfluidic device may be coupled with
downstream
operations such as secondary microfluidic dissociation devices to better
liberate single cells
from small aggregates. Valves may also optionally be incorporated into or
associated with the
upstream hydro-mincing microfluidic channels to provide a high degree of shear
forces on
selected or targeted areas or regions of tissue. These valves can be turned on
and off to cover
the entire length of tissue in the chamber. In addition, cell sorting and
analysis components
may be added to create point-of-care platforms for cell-based diagnostics and
therapies.
[0008] In another embodiment, a method of processing tissue using the
microfluidic
device includes placing the tissue within the sample chamber and then flowing
a fluid
containing a digestive enzyme into the inlet. The tissue that may be processed
using the
microfluidic device may include healthy or diseased tissue. For example, in
one particular
embodiment, the tissue that is processed by the device includes tumor tissue,
although other
tissue types are contemplated. Tissue obtained from different organs may also
be treated.
Examples include liver tissue, kidney tissue, pancreas tissue, spleen tissue,
skin tissue, heart
tissue, and the like. The fluid may be pumped into the microfluidic device
using a pump. The
cells or smaller aggregates of tissue may be collected from the outlet of the
microfluidic
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device. In some embodiments, the collected output from the microfluidic device
is
recirculated back into the input of the microfluidic device.
[0009] In some embodiments, the tissue is loaded into the sample chamber by
using a
sample port. In some embodiments the sample is loaded by inserting a needle
into the sample
port and depositing the tissue in the sample chamber. In other embodiments, a
plug, cap, or
lid covers the sample chamber and can be removed/secured to the microfluidic
device.
[0010] A microfluidic system for the processing of a tissue sample
dimensioned within the
range of 1 mm3 to 50 mm3 into cellular suspensions including a microfluidic
device formed
from a substrate or chip having formed therein an inlet, an outlet, and a
sample chamber
dimensioned to hold the tissue sample, the sample chamber fluidically coupled
at a first side
to a plurality of upstream hydro-mincing microfluidic channels disposed in the
substrate or
chip further fluidically coupled to the inlet and coupled at a second side of
the sample
chamber to a plurality of downstream sieve microfluidic channels disposed in
the substrate or
chip further fluidically coupled to the outlet; wherein both the width of the
upstream hydro-
mincing microfluidic and the width of the downstream sieve microfluidic
channels are greater
than 50 pm and are smaller than the smallest dimension of the tissue sample.
[0011] A method of processing tissue in a microfluidic device that is
formed in a substrate
or chip having formed therein an inlet, an outlet, and a sample chamber
dimensioned to hold
the tissue sample, the sample chamber fluidically coupled at one side to a
plurality of
upstream hydro-mincing microfluidic channels disposed in the substrate or chip
and further
fluidically coupled to the inlet and coupled at another side of the sample
chamber to a
plurality of downstream sieve microfluidic channels disposed in the substrate
or chip and
further fluidically coupled to the outlet. The method includes placing the
tissue within the
sample chamber and flowing a fluid containing a digestive enzyme into the
inlet.
Brief Description of the Drawin2s
[0012] The features and advantages of the present invention will become
apparent from a
consideration of the following detailed description presented in connection
with the
accompanying drawings in which:
[0013] FIG. 1 illustrates a top view of one embodiment of a microfluidic
device for the
processing and digestion of tissue according to one embodiment.
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[0014] FIG. 2 illustrates a schematic illustration of a system that
incorporates a
microfluidic device for the processing and digestion of tissue with a
secondary microfluidic
device that is located downstream of the first device.
[0015] FIG. 3 illustrates a photographic image of a laser-etched acrylic
sheet containing
the chamber for loading tissue samples and fluidic "mincing" channels
including upstream
(left) for hydro-mincing and downstream (right) sieves (sieve gates).
[0016] FIG. 4 illustrates an exploded view of a microfluidic device
according to one
embodiment that includes a polymer or elastic gasket layer sandwiched between
two acrylic
sheets or other hard plastic sheets. Hose barbs are illustrated in the top
layer and nylon screws
are used to hold the device together.
[0017] FIG. 5 illustrates a photograph of the fully assembled device
illustrated in FIG. 4.
[0018] FIG. 6 schematically illustrates the experimental set-up used for
digestion
experiments using the microfluidic device for the processing and digestion of
tissue. Flow
was driven by a peristaltic pump and tissue digestion was visually monitored
with a camera
mounted above the device.
[0019] FIG. 7 illustrates finite-element fluid dynamics simulations showing
velocity
profiles in devices with different numbers of hydro-mincing microfluidic
channels (3, 5, and
7). Simulation results are shown at 1 mL/min flow rate with the chamber empty
and partially
blocked by a model tissue. Fewer hydro-mince channels will generate stronger
fluidic jets to
shear the tissue, but with less overall coverage.
[0020] FIG. 8A illustrates an exploded view of another embodiment of a
microfluidic
device for the processing and digestion of tissue according to one embodiment.
[0021] FIG. 8B illustrates a top or plan view of the microfluidic device of
FIG. 8A.
[0022] FIG. 9A illustrates an exploded view of another embodiment of a
microfluidic
device for the processing and digestion of tissue according to one embodiment.
[0023] FIG. 9B illustrates a top or plan view of the microfluidic device of
FIG. 9A.
[0024] FIG. 10 illustrates another embodiment of a microfluidic device that
includes
valves located in the hydro-mincing microfluidic channels.
[0025] FIG. 11 illustrates an exploded view of another embodiment of a
microfluidic
device for the processing and digestion of tissue according to one embodiment.
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[0026] FIG. 12 illustrates an exploded view of another embodiment of a
microfluidic
device for the processing and digestion of tissue according to one embodiment.
[0027] FIG. 13A is a photographic image of a tissue core obtained using a
Tru-CutTm
biopsy needle and placed inside the tissue chamber.
[0028] FIG. 13B illustrates time-lapse images of tissue digestion for
devices with 3, 5, and
7 hydro-mincing microfluidic channels. The fluid contained collagenase, and
was pumped
through the device at 20 mL/min.
[0029] FIG. 13C illustrates a graph showing tissue loss as a function of
time. Tissue loss
was quantified from images based on mean gray value and overall tissue area.
Trends were
similar for each design, but variability was lowest for 3 hydro-mincing
microfluidic channels.
[0030] FIG. 13D illustrates micrograph images of device effluents after 30
min operation.
Top is seven (7) hydro-mincing microfluidic channels. Middle is five (5) hydro-
mincing
microfluidic channels. Bottom is three (3) is hydro-mincing microfluidic
channels. Scale bar
is 100 um. Error bars represent standard errors from at least three
independent experiments.
[0031] FIG. 14 illustrates an image processing algorithm used to monitor
tissue digestion.
Images were analyzed for tissue size and density to quantify changes during
digestion within
the device. First, raw images were separately converted to binary (upper
arrow) and grayscale
(lower arrow) images to outline the contour and quantify mean gray value,
respectively. The
area within the tissue contour was then calculated, and multiplied by mean
gray value to
obtain a single metric accounting for tissue size and density.
[0032] FIG. 15A illustrates photographic images of mouse liver (top right)
and kidneys
(bottom right) were freshly harvested and cut into 1 cm long x 1 mm diameter
pieces and
placed within the device sample chamber.
[0033] FIG. 15B illustrate time-lapsed images of tissue digestion for
devices containing 3
hydro-mincing microfluidic channels. Tissue size and density both decreased
over time as
digestion progressed.
[0034] FIG. 15C illustrates a graph of tissue loss quantified from images
based on mean
gray value and overall tissue area, with liver and kidney samples
demonstrating similar
trends.
[0035] FIG. 15D illustrates a graph illustrating the results of the
CyQUANTO assay was
used to directly quantify cell suspensions obtained by digestion only, scalpel
mincing and
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digestion, or device treatment lasting for a total of 15, 30, or 60 min. The
CyQUANTO signal
increased with treatment time, and was higher overall for kidney samples.
Signals from
device treated samples were consistently higher than minced controls, similar
to gDNA and
cell counting results presented in FIG. 3 of the main text. Error bars
represent standard errors
from at least three independent experiments. * indicates p < 0.05 relative to
minced control at
the same digestion time.
[0036] FIG. 16A illustrates a graph showing the amount of genomic DNA (gDNA)
that
was extracted and quantified from kidney and liver tissue cell suspensions
obtained by
digestion only, scalpel mincing and digestion, or device treatment lasting for
a total of 15, 30,
or 60 min. As seen in the graph, gDNA increased with treatment time, and
overall was higher
for kidney samples. Device treatment consistently provided more gDNA than
minced
controls at the same time point. In most cases, gDNA was also higher than the
next digestion
time point, although differences were not significant.
[0037] FIG. 16B illustrates a graph showing cell counter results, showing
that single cell
numbers largely matched gDNA findings but with higher variability. Also, liver
values were
now similarly comparable to kidney, suggesting that kidney suspensions may
have contained
more aggregates. Error bars represent standard errors from at least three
independent
experiments. * indicates p < 0.05 relative to minced control at the same
digestion time.
[0038] FIG. 16C illustrates micrographs of minced controls and device
effluents after
lysing red blood cells. Note the large number of aggregates in the controls,
particularly at 60
min. Scale bar is 100 pm.
[0039] FIG. 17 illustrates FACS gating data for cell suspensions obtained
from digested
mouse and liver and kidney samples. Cell suspensions obtained from digested
mouse liver
and kidney samples were stained with the four-probe panel listed in Table 1
and analyzed
using flow cytometry. Controls were treated only with an isotype matched
(IgG2b), PE-
conjugated antibody. Acquired data was assessed using a sequential gating
scheme. First, an
FSC-A vs. SSC-A gate (Gate 1) was used to exclude debris near the origin. Gate
2 was based
on FSC-A vs. FSC-H, and was used to select single cells. Gate 3 distinguished
CD45+
leukocytes based on CD45-PE signals in FL2-A vs. SSC-H plots. The CD45- cell
subset was
further divided into anucleate RBCs and nucleated tissue cell subsets based on
signals from
the Draq5 nuclear stain in FL4-A vs. SSC-H plots. The cellularity of nucleated
tissue cells of
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interest was validated based on the signal of the cell membrane dye CellMaskTm
Green in
FLI-A vs. FSC-H plots. Finally, live and dead tissue cells were discriminated
based on 7AAD
signals in FL3-A vs. SSC-H plots. All gates were established using the minced
control that
was digested for 60 min. Heat treated cells were used as a positive control to
confirm
appropriate 7AAD signals for dead cells.
[0040] FIG. 18A illustrates flow cytometry results of mouse kidney
suspensions. Flow
cytometry was used to identify and quantify the number of leukocytes, red
blood cells, and
single tissue cells in the suspensions obtained from minced controls or device
treatment.
Relative numbers of each cell type are shown. Red blood cells comprised the
highest
percentage of almost all populations, and there was no statistically
significant change in
population compositions across all minced control and device conditions.
[0041] FIG. 18B illustrates flow cytometry results of mouse liver
suspensions. Flow
cytometry was used to identify and quantify the number of leukocytes, red
blood cells, and
single tissue cells in the suspensions obtained from minced controls or device
treatment.
Relative numbers of each cell type are shown. Red blood cells comprised the
highest
percentage of almost all populations, and there was no statistically
significant change in
population compositions across all minced control and device conditions.
[0042] FIG. 18C illustrates a graph of total and live tissue cell numbers
per mg of tissue
for kidney samples.
[0043] FIG. 18D illustrates a graph of total and live tissue cell numbers
per mg of tissue
for liver samples. For both FIG. 18C and 18D, tissue cell recovery increased
with digestion
time for minced controls, but did not change significantly with device
processing beyond 10
min. Importantly though, all device conditions yielded more cells than minced
controls that
were digested for up to 30 min. Viability remained >80% for all but the
longest time points,
which reached as low as 70%. The x-axis for FIGS. 18A and 18B is the same as
FIGS. 18C
and 18D. Error bars represent standard errors from at least three independent
experiments. *
indicates p < 0.05 relative to minced control at the same digestion time. #
indicates p < 0.05
compared to minced control digested for 15 min.
[0044] FIG. 19 illustrates a graph illustrating mouse kidney and liver cell
viability data.
Cell viability was similar for device treated conditions relative to minced
counterparts,
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demonstrating minimal effect of device treatment. Error bars represent
standard errors from
at least three independent experiments.
[0045] FIG. 20 shows an embodiment of the microfluidic device of the
present invention
having an equal number of upstream hydro-mincing microfluidic channels and
downstream
sieve microfluidic channels, causing the device to be symmetrical.
[0046] FIG. 21A shows a photograph of a plurality of microfluidic devices
coupled in
parallel at the sample chamber in order to process larger sample sizes.
[0047] FIG. 21B shows a schematic of a plurality of microfluidic devices
coupled in
parallel at the sample chamber in order to process larger sample sizes.
[0048] FIG. 22 shows a schematic of an embodiment of the present invention
wherein the
outlet is fluidly connected to a junction for directing pieces of digested
tissue sample through
an exit tube out of the system or recirculated back into the inlet of the
device in order to
further process the tissue sample. A digestive enzyme source is additionally
fluidly connected
to a digestive enzyme source for providing additional digestive enzyme to be
introduced to
the inlet of the microfluidic device to replace digested tissue samples that
exit the system
through the outlet.
Detailed Description of the Illustrated Embodiments
[0049] FIG. 1 illustrates microfluidic device 10 for the processing and
digestion of tissue
according to one embodiment. In one particular embodiment, the microfluidic
device 10 is
designed to process tissue obtained from a mammalian subject (e.g., human). In
particular,
the microfluidic device 10 has particular applicability for the processing and
digestion of
tumor tissues, although other tissue types may be processed using the
microfluidic device 10.
The microfluidic device 10 is formed in a substrate or chip structure 12 which
as explained
herein may be formed using multiple layers that are assembled together to form
the
microfluidic device 10. The microfluidic device 10 includes three primary
features that are
defined or formed in the substrate or chip structure 12. These include a
sample chamber 14
that holds a sample 16 in place while fluid containing proteolytic enzymes or
other digestive
agents is passed into the sample chamber 14 and onto the surface of the sample
16. The
sample chamber 14 thus maintains the sample 16 in a generally fixed location
in the substrate
or chip structure 12. By having the sample 16 be retained in the sample
chamber 14, this
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promotes sample mixing, enhances enzymatic activity, and as explained below
applies
hydrodynamic shear forces to mechanically dislodge cells and aggregates from
the larger
sample 16.
[0050] In some embodiments, the present invention features a system for the
processing of
a sample 16 into cellular suspensions. The system may include a tissue sample
16 having a
size within the range of 1 mm3 to 50 mm3. The system may further comprise a
microfluidic
device 10. The microfluidic device 10 is formed as a substrate or chip 12. The
substrate or
chip 12 may have an inlet 22, an outlet 28, and a sample chamber 14
dimensioned to hold the
tissue sample 16 formed therein. The sample chamber 14 may be fluidically
coupled at a first
side to a plurality of upstream hydro-mincing microfluidic channels 18
disposed in the
substrate or chip 12. The upstream hydro-mincing microfluidic channels 18 may
be further
fluidically coupled to the inlet 22. The sample chamber 14 may additionally be
coupled at a
second side to a plurality of downstream sieve microfluidic channels 24
disposed in the
substrate or chip 12. The downstream sieve microfluidic channels 24 may be
further
fluidically coupled to the outlet 28. In some embodiments, the width of the
upstream hydro-
mincing microfluidic channels 18 and the width of the downstream sieve
microfluidic
channels 24 may be greater than 50 nm. In some embodiments, the width of the
upstream
hydro-mincing microfluidic channels 18 and the width of the downstream sieve
microfluidic
channels 24 may be smaller than the smallest dimension of the tissue sample
16. In some
embodiments, the width of the sample chamber 14 may be within the range of
about 0.5 mm
and 1 cm. In some embodiments, the length of the sample chamber 14 may be less
than 50
cm, and the height of the sample chamber 14 may be less than 5 cm. In some
embodiments,
the number of upstream hydro-mincing microfluidic channels 18 may be equal to
the number
of downstream sieve microfluidic channels 24 (see FIG. 20). The number of
upstream hydro-
mincing microfluidic channels 18 and downstream sieve microfluidic channels 24
and the
width of the sample chamber 14 may depend on the size of the tissue sample 16
being
processed. Tissue samples 16 with a smaller width may be processed more
effectively than
tissue samples 16 with a larger width.
[0051] In some embodiments, a first instance of the microfluidic device 10
may be
coupled to at most two additional instances of the microfluidic device 10. The
coupling may
occur at a third side of the sample chamber 14 (e.g., a side), a fourth side
of the sample
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chamber 14 (e.g., another side), or a combination thereof depending on how
many instances
of the microfluidic device 10 are coupled to the first instance of the
microfluidic device 10.
These different instances may be contained in the same substrate or chip 12 as
seen in FIGS.
21A and 12B. FIGS. 21A-21B show embodiments of the aforementioned parallel
coupling
of microfluidic devices 10. This allows for an indefinite number of
microfluidic devices 10 to
be coupled to each other in parallel in order to process larger sample sizes
without the need to
lengthen or enlarge the device. The flow rate of fluid directed through the
coupled
microfluidic devices 10 may be proportionally increased depending on the
number of
instances of microfluidic devices 10 coupled together. In some embodiments, a
different type
of tissue sample 16 is placed in the sample chamber 14 of each instance of the
microfluidic
device. The type of tissue sample 16 processed by the present invention may be
selected from
a group comprising kidney tissue, liver tissue, heart tissue, lung tissue,
breast tumor tissue,
spleen tissue, and pancreas tissue.
[0052] The microfluidic device 10, in one embodiment, was designed to
process samples
obtained from core needle biopsies, directly into cell suspensions without the
need for manual
processing steps such as scalpel mincing. However, in other embodiments, the
microfluidic
device 10 processes a larger tissue sample after the sample has been subject
to some
mechanical processing (e.g., scalpel mincing). The particular size of the
sample chamber 14
may vary depending on the size of the sample 16. Typically, the width of the
sample chamber
14 may be within the range of about 0.5 mm and about 10 mm, the length of the
sample
chamber 14 is less than 2 cm, and the height of the sample chamber 14 is less
than 1 cm. For
example, with reference to FIG. 1, in one embodiment, the sample chamber 14
has a width 2
mm or less, the length of the sample chamber 14 is less than 2 cm, and the
height of the
sample chamber 14 is less than 2 mm (height dimension is perpendicular to the
plane of the
page). Fluid flows in the direction of arrows A along the width of the sample
chamber 14. In
another embodiment, the width of the sample chamber 14 is around 1.5 mm or
less, the
length of the sample chamber 14 is around 1 cm or less (e.g., approximately
the size of a Tru-
Cut core biopsy needle; about 1 cm long x 1 mm diameter tissue), and the
height of the
chamber is less than 2 mm (e.g., around 1 mm). In still other embodiments, the
sample
chamber 14 may be much smaller, for example, and can accommodate samples 16
having a
longest dimension of around 1 mm. For example, when the sample 16 has been
subject to
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some mechanical processing (e.g., mincing) the sample chamber 14 may have a
much smaller
size. The height of the sample chamber 14 may vary but is typically less than
1 cm and more
typically less than 2 mm (e.g., a height of 1 mm may be used as described
herein). In some
embodiments, the sample chamber 14 is dimensioned to receive a sample 16
directly obtained
from a tissue biopsy device such as the Tru-Cut core biopsy needle or similar
devices.
[0053] The second feature of the microfluidic device 10 includes a
plurality of hydro-
mincing microfluidic channels 18 located upstream of the sample chamber 14
which focus
fluid into high velocity jets that are directed into the sample 16 retained in
the sample
chamber 14. The hydro-mincing microfluidic channels 18 as seen in FIG. 1 are
fluidically
coupled with an inlet channel 20 that receives fluid from an inlet 22. Fluid
thus moves
through the microfluidic device 10 in the direction of arrows A. The hydro-
mincing
microfluidic channels 18 produce fluid jets that concentrate hydrodynamic
shear forces at
discrete locations on the sample 16, breaking the sample 16 down mechanically
and
delivering proteolytic enzymes deep inside the sample 16 (i.e., tissue). This
is analogous to
manually mincing the tissue with a scalpel, and hence these are referred to as
hydro-mincing
microfluidic channels 18. The width of the hydro-mincing microfluidic channels
18 may vary
but is generally within the range of about 50 um to about 1 mm. For example, a
width of the
hydro-mincing microfluidic channels 18 within the range of about 100 um to
about 200 um
may be typical, although other dimensions outside this specific range may be
used.
[0054] Finally, a plurality of downstream sieve microfluidic channels 24
are located
downstream of the sample chamber 14 to act as a sieve that selectively retains
larger pieces of
tissue and cellular aggregates for further digestion. The downstream sieve
microfluidic
channels 24 form sieve gates that retain the larger sized tissue portions and
cellular
aggregates to prevent them from passing further downstream. Smaller aggregates
and single
cells are, however, allowed to pass out of the device 10 for collection or
potentially for
further microfluidic processing. For example, the cells that leave the device
10 may be
subject to downstream cell sorting and/or analysis to create point-of-care
platforms for cell-
based diagnostics and therapies.
[0055] In one embodiment, the downstream sieve microfluidic channels 24
(along with
the hydro-mincing microfluidic channels 18) are spaced evenly along the side
or end of the
sample chamber 14 to firmly secure the sample 16 in place in the sample
chamber 14 and
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minimize backpressure. The width of the downstream sieve microfluidic channels
24 may
vary but may be within the range of about 10 p.m and 1 mm. More typically, the
downstream
sieve microfluidic channels 24 have a width within the range of about 100 p.m
to about 1 mm.
For example, in some embodiments a width within the range of 500 p.m to lmm is
useful.
During experiments described herein, a channel width of 500 p.m for the
downstream sieve
microfluidic channels 24 was used and the device could comfortably accommodate
seven (7)
such channels across the width of the sample chamber 16. The plurality of
downstream sieve
microfluidic channels 24 lead to a common outlet channel 26 that extends to an
outlet 28
where fluid can leave the microfluidic device 10.
[0056] In some embodiments, the width of the downstream sieve microfluidic
channels 24
may be larger than the width of the hydro-mincing microfluidic channels 18. In
other
embodiments, the width of the downstream sieve microfluidic channels 24 may be
smaller
than the width of the hydro-mincing microfluidic channels 18. In still other
embodiments, the
width of the downstream sieve microfluidic channels 24 may be substantially
the same as the
width of the hydro-mincing microfluidic channels 18.
[0057] Note that 500 p.m is comparable to the ¨1 mm size of tissue pieces
typically
achieved by scalpel mincing. Aggregates of this size would also be ideal for
directly inputting
into a downstream branching channel array dissociation device such as that
disclosed in U.S.
Patent No. 9,580,678, which is incorporated therein. FIG. 2 illustrates one
such embodiment
where the output from a first device (i.e., Device#1; microfluidic device 10)
is then input into
a second downstream device 100 (Device #2) for additional tissue dissociation.
[0058] For example, in one embodiment, the microfluidic device 10 is
coupled to another
tissue dissociation device 100 like that illustrated in the '678 patent. In
that device 100, a
series of stages of microfluidic channels with decreasing dimensions and
having a series of
expansion and constriction regions (illustrated in FIG. 2) are used to impart
shearing forces
on cell clusters and aggregates to dissociate tissue. The microfluidic device
10 can be coupled
to such as device 100 as is illustrated in FIG. 2. In this embodiment, the
output of the first
microfluidic device 10 may be coupled to the input of the second microfluidic
device 100 that
is used for further tissue dissociation. A valve 30 may be located between the
two devices
(10, 100) to allow selective flow through the second tissue dissociation
device 100. In
addition, a pump 32 is illustrated that is used to recirculate flow between
one or both of the
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microfluidic devices 10, 100. It should be understood, however, that rather
than recirculating
flow as illustrated, flow may pass directly through the devices without any
recirculation.
[0059] In some embodiments, the outlet 28 of the microfluidic device 10 may
be fluidly
connected to a junction. The junction may be fluidly connected to both an exit
tube 31 and a
recirculation tube 33. The exit tube 31 may be configured such that the tissue
sample 16 may
be directed by the pump 32 through the exit tube 31 to a collection chamber.
The
recirculation tube 33 may be fluidly connected to the inlet 22 of the
microfluidic device 10
and configured such that the digestive enzyme fluid directed through the
microfluidic device
may be directed by the pump 32 through the recirculation tube 33 to the inlet
22. In some
embodiments, the recirculation tube 33 may additionally be fluidly connected
to a digestive
enzyme source for providing additional digestive enzyme to be introduced into
the
microfluidic device 10 (see FIG. 22). This process is used to further process
the tissue sample
16.
[0060] For the plurality of hydro-mincing microfluidic channels 18, the
goal is to achieve
efficient hydro-mincing. Using fewer channels would generate stronger fluidic
jets, but
would also cover less of the tissue cross-section and lead to higher device
back-pressures.
Since these are competing factors, experiments were conducted to use channel
number as a
test variable and created devices with three (3), five (5), and seven (7)
hydro-mincing
microfluidic channels 18. FIG. 3 illustrates a photograph of the microfluidic
device 10
formed in hard acrylic sheets showing three (3) hydro-mincing microfluidic
channels 18 and
seven (7) downstream sieve microfluidic channels 24. Alternatively, materials
for the
microfluidic device 10 include polyethylene terephthalate (PET). FIG. 4
illustrates the multi-
layered construction of the microfluidic device 10 according to one
embodiment. In this
embodiment, the microfluidic device 10 an upper substrate layer 12a formed
from PET that
includes the barbed ends or tubing connections 34 (e.g., hose barbs) that are
fluidically
coupled to the inlet 22 and outlet 28. The microfluidic device 10 further
includes a lower
substrate 12b that has the microfluidic features formed therein. This includes
the inlet 22,
inlet channel 20, sample chamber 14, downstream sieve microfluidic channels
24, outlet
channel 26, and outlet 28 of FIG. 1. In this particular embodiment a
polydimethylsiloxane
(PDMS) gasket 12c having holes or vias 35 (for fluid access) is sandwiched
between the
upper substrate 12a and the lower substrate 12b which collectively together
form the
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substrate/chip 12. Other elastic or polymers may be used for the gasket 12c.
In this particular
embodiment, fasteners 36 (e.g., screws) were used to secure the multiple
substrates 12a, 12b,
12c together in a single construction as seen in FIG. 5.
[0061] As for channel size, smaller widths would generate stronger, more
concentrated
fluidic jets. Therefore, for experiments conducted on the microfluidic device
10 a width of
200 p.m was chosen for the hydro-mincing microfluidic channels 18, which was
the smallest
feature resolution that could reliably be achieved with the laser-based
fabrication method. It
should be understood, however, that other dimensions may be used for the
upstream hydro-
mincing microfluidic channels 18 as well as the downstream sieve microfluidic
channels 24.
[0062] Devices 10 were fabricated in hard acrylic sheets using a laser to
etch the sample
chamber 14 and channel features of the hydro-mincing microfluidic channels 18,
downstream
sieve microfluidic channels 24, inlet channel 20, inlet 22, outlet channel 26,
and outlet 28 in a
first substrate 12a as described above. Laser power and raster speed were
controlled to
achieve a depth of approximately 1 mm, establishing channel height. A second
layer of
acrylic was used as the second substrate 12b and was tapped and fitted with
hose barbs 34 to
connect inlet and outlet tubing. Finally, the gasket layer 12c composed of
polydimethylsiloxane (PDMS) was sandwiched between the acrylic layers 12a, 12b
to
provide a watertight seal. Note that the deformable nature of PDMS, and likely
the tissue
itself, should alleviate fluid flow and backpressure issues even while the
tissue is initially
obstructing the flow path. Finally, the assembled device sandwich 10 was held
together using
six (6) nylon screws 36 as seen in FIG. 5. The experimental set-up is shown in
FIG. 6. For
initial experiments, a peristaltic pump 32 was used to recirculate fluid
through the device to
conserve proteolytic enzyme solution. Of course, in an alternative embodiment,
the flow may
be continuous or the cells removed prior to recirculation in the device. In
the experimental
apparatus, a camera 38 was mounted above the microfluidic device 10 to monitor
the
progress of tissue digestion.
[0063] Computational fluid dynamics simulations were performed using COMSOL
Multiphysics software for each three (3), five (5), and seven (7) hydro-
mincing microfluidic
channels 18 using a flow rate of 1 mL/min (FIG. 7). These simulations were
performed with
and without a model tissue within the sample chamber 14 to obstruct flow. As
expected, the
design with three (3) hydro-mincing microfluidic channels 18 generated the
highest fluid
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velocities, or strongest fluidic "cuts." Increasing the number of hydro-
mincing microfluidic
channels 18 provided weaker "cuts" that were better dispersed across the
tissue.
[0064] FIGS. 8A, 8B, 9A, and 9B illustrate two alternative embodiments of a
microfluidic
device 10 that includes a feature to aid in loading the tissue or other sample
into the sample
chamber 16. The microfluidic device 10 includes a plurality of layers 12a,
12b, 12c that may
be pressure laminated together with the aid of an adhesive or glue applied to
the interface
between adjacent layers to form the microfluidic device 10. Alternatively, the
layers 12a,
12b, 12c may be secured together using fasteners such as those used in the
embodiments of
FIGS. 4 and 4. In the embodiment of FIGS. 8A and 8B, an open window 40 is
provided on
top layer 12a of the microfluidic device 10 through which sample 16 can be
loaded using
forceps. A plug 42 that is formed from silicon rubber material or the like is
then placed in the
window and secured with adhesive tape or glue. The middle layer 12c includes a
hole or via
44 formed therein so that the sample 16 can be loaded into the sample chamber
14 formed in
the first substrate 12a.
[0065] In the embodiment of FIGS. 9A and 9B, the microfluidic device 10 has
a sample
port 46 located in the side of the microfluidic device 10 that can be used to
load a sample 16
into the sample chamber 14. This design enables access through the side of the
microfluidic
device 10 by penetrating a silicon rubber septum 48 with a needle 50 as seen
in FIG. 9B. In
one embodiment, this needle 50 would be the same one used to extract the
sample 16 (e.g.,
tissue), such as a core needle or punch biopsy. After penetrating the septum
48, the needle 50
would seat the sample 16 in the sample chamber 14 and remain in place
(penetrating the
septum 48) to seal the microfluidic device 10. In an alternative embodiment,
the septum 48
may be a self-sealing septum 48 such that the needle 50 can be removed from
the
microfluidic device 10 without leakage of fluid or other contents outside of
the microfluidic
device 10.
[0066] FIG. 10 illustrates one alternative embodiment of a microfluidic
device 10 where
one or more of the hydro-mincing microfluidic channels 18 can be selectively
turned on or
off by way of individual valves 54. The direction of fluid flow is indicated
by arrow A in
FIG. 10. In this regard, the option is provided to mince a large area of the
sample 16 by
sequentially using a small number of hydro-mincing microfluidic channels 18.
For example,
one or few of the hydro-mincing microfluidic channels 18 may be opened at any
particular
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time so that high jetting or shearing forces are imparted on the sample 16.
The valve(s) 54
may then be closed and another valve 54 or set of valves 54 that are aimed at
a different
region of sample 16 can then be turned on. The valves 54 may include
microfluidic valves 54
that are known in the art. For example, microfluidic valves 54 that use a
deformable
membrane to actuate flow in a channel are known and may be used as one
example. This
process may continue for any number of cycles to mince the entire sample 16.
100671 FIG. 11 illustrates another embodiment of the microfluidic device
10. This
microfluidic device 10 is formed from a multi-layer construction using hard
acrylic or PET as
described herein. In this embodiment, the microfluidic device 10 is formed
from multiple
layers 60a, 60b, 60c, 60d, 60e, and 60f The layers 60a, 60b, 60c, 60d, 60e,
and 60f are
pressure laminated together with the aid of an adhesive or glue (e.g.,
silicone or acrylic-based
glues or adhesives) applied to the interface between adjacent layers to form
the microfluidic
device 10. Layer 60a serves as a base or bottom layer. Layer 60b has formed
therein the inlet
channels 20, outlet channels 26, as well as the hydro-mincing microfluidic
channels 18 and
downstream sieve microfluidic channels 24. Layer 60c has formed therein vias
62 that
communicate with the inlet channels 20 and outlet channels 26 in layer 60b as
well as
apertures or holes 64 that provide access for the inlet 22 and the outlet 28.
Layer 60d includes
the sample chamber 14 formed therein that communicates with the vias 62 in
layer 60c. Layer
60d further includes apertures or holes 64. Layer 60e includes in addition to
the apertures or
holes 64 a via 66 that provides access to the sample chamber 14. This via 66
may have a
diameter of around 1 mm which is sized to accommodate sample 16 that has been
mechanically processed (e.g., minced). Layer 60f is the top layer and includes
barbed ends 34
that provide fluid access in/out of the microfluidic device 10. In addition,
the layer 60f
includes a loading port 68 that communicates with the via 66 and into the
sample chamber
14. A cap (not shown) may be placed over the loading port 68 after loading so
that fluid and
tissue/cells remain inside the microfluidic device 10.
100681 The loading port 68 may be configured as a Luer end that interfaces
with a syringe
or the like for loading. In this embodiment, minced sample 16 (e.g., minced
tissue) is loaded
into the loading port 68 prior to flowing fluid through the microfluidic
device. In one
alternative of this embodiment, the downstream sieve microfluidic channels 24
that
communicate with the outlet channels 26 may include a filtering capability
that restrict the
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passage of larger pieces of sample 16 from flowing downstream in the device
10. Filtering
may also be provided by the vias 62.
[0069] FIG. 12 illustrates another embodiment of the microfluidic device
10. This
microfluidic device 10 is formed from a multi-layer construction using hard
acrylic or PET as
described herein. In this embodiment, the microfluidic device 10 is formed
from multiple
layers 70a, 70b, 70c, 70d and includes a cap or lid 72 that, as explained
herein, is used to
close the device after the sample 16 has been loaded into the microfluidic
device 10. The
layers 70a, 70b, 70c, 70d are pressure laminated together with the aid of an
adhesive or glue
applied to the interface between adjacent layers to form the microfluidic
device 10. Layer 70a
serves as a base or bottom layer. Layer 70b has formed therein the inlet
channels 20 and
outlet channels 26. Layer 70c has formed therein vias 74 that communicate with
the inlet
channels 20 and outlet channels 26 in layer 70b as well as apertures or holes
76 that provide
access for the inlet 22 and the outlet 28. Layer 70d includes the sample
chamber 14 formed
therein that communicates with the vias 74 in layer 70c. Layer 70d also
includes apertures or
holes 78 that can accommodate barbed ends 34 (not illustrated in FIG. 12) such
as those
illustrated in FIGS. 4, 5, 8A, 8B, 9A, 9B, and 11.
[0070] In this embodiment, the sample 16 can be loaded directly into the
sample chamber
14. After loading the sample 16 in the sample chamber 14, the cap or lid 72 is
then affixed to
the layer 70d above the sample chamber 14 to seal the sample chamber 14 from
the external
environment of the microfluidic device 10. The cap or lid 72 may be secured to
the layer 70d
using an adhesive or the like. In some embodiments, the cap or like 72 may be
removable so
that the microfluidic device 10 can be used multiple times. In other
embodiments, however,
the cap or lid 72 is secured to the layer 70d in a permanent manner. As an
alternative to an
adhesive or glue, the cap or lid 72 may be secured to the layer 70d using one
or more
fasteners (not shown) such as clamps, screws, bands, clips, or the like.
[0071] Initial Device Optimization Using Beef Liver Tissue. Performance of
the
microfluidic digestion device such as that illustrated in FIG. 1 was first
evaluated using beef
liver as the sample tissue. Model tissue cores were extracted using a Tru-Cut
biopsy needle
and loaded into the sample chamber 14 (FIG. 13A). Devices were then primed
with PBS
buffers containing collagenase, sealed, and flow was initiated at 20 mL/min,
the highest flow
rate that could be achieved with the peristaltic pump. Images of tissue
specimens were
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acquired every 5 min using the camera 38 mounted above the device as seen in
FIG. 6 to
monitor digestion, and experiments were performed for a total of 30 min (FIG.
13B). After
each image was acquired, flow was briefly reversed to clear tissue that had
seeped into the
sieve microfluidic channels. Tissue seeping was most extensive using three (3)
hydro-mince
channels, reflecting the higher hydrodynamic forces being generated. Images
were processed
using ImageJ and MATLAB to assess the amount of liver tissue remaining in the
device at
each time point based on tissue area and pixel density (see FIG. 14).
Digestion profiles are
plotted in FIG. 13C, after normalizing by initial tissue mass. Results were
nominally similar
for all three devices, with a dramatic 40% tissue decrease during the first 5
min, followed by
a more gradual decrease in tissue by ¨10% per 5 min interval. The initial drop
primarily
correlated to diminished pixel density, which may have reflected tissue
debulking or washing
out of red blood cells. The subsequent gradual phase primarily reflected a
loss of tissue mass.
After 30 min, approximately 80% of the tissue had been removed from all three
of the device
designs. However, the device with three (3) hydro-mincing microfluidic
channels provided
the most consistent results in terms of lower variability between experiments,
particularly at
later time points, and thus was chosen for further evaluation. Representative
micrographs of
device effluents collected after 30 min device processing are shown in FIG.
13D. For all
cases, sample effluents primarily comprised a mixture of larger tissue
aggregates, tissue cells,
and red blood cells.
[0072] Evaluation of Cell Suspensions Obtained from Fresh Mouse Organs. Next,
the
three (3) hydro-mincing channel design was tested using freshly resected
murine liver and
kidney samples. These live tissues better represent samples that will be used
in future
applications, and the resulting cell suspensions can be directly assayed for
quality. Liver is
generally considered to be among the easiest tissues to dissociate, but
hepatocytes are well
known to be fragile. Kidney is considered to be a difficult tissue to
dissociate due to its
structure as a dense array of blood vessels and epithelial lined tubules,
which function under
high physiologic hydrodynamic pressures, have tight intercellular junctions,
and have
specialized basement membranes. Immediately after harvesting, tissues were cut
into ¨1 cm x
1 mm x 1 mm pieces with a scalpel (see FIG. 15A) and weighed. Digestion device
experiments were then conducted as described for beef liver, with collagenase
recirculated
for either 15 or 30 min before sample collection. Images were again taken
every 5 min and
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processed to monitor tissue loss, which was similar to beef liver (see FIGS.
15B and 15C).
Controls were further minced with a scalpel into ¨1 mm3 pieces before
digesting with
collagenase for 15, 30, or 60 min in a conical tube. These samples were
constantly agitated,
and vortexed every 5 min. A separate control was included in which the tissue
was not
minced, only digested for 30 min. Following digestion, device-processed and
control samples
were mechanically treated by vortexing and pipetting, filtered through a 70 pm
cell strainer,
and treated with DNase to remove extracellular DNA. Cellular content was then
assessed
based on total genomic DNA (gDNA) extracted using a QIAamp0 DNA kit. For
minced
controls, gDNA progressively increased with digestion time (FIG. 16A). Kidney
samples
yielded approximately 100 ng gDNA per mg of tissue after 60 min digestion,
while liver was
less than half this value. Slightly less gDNA was obtained from the un-minced
controls, but
differences were not significant. Device treatment yielded dramatically more
gDNA than
controls when compared at the same digestion time. The difference was
approximately 5-fold
for both tissue types after 15 min, and 3 to 4-fold after 30 min. Moreover,
device treatment
produced at least as much gDNA as the minced control at the next longer
digestion time
point. Thus, the microfluidic digestion device can significantly improve
digestion efficiency
and shorten digestion time. DNA was also assessed within intact cellular
suspensions using
the CyQUANTO assay, which corroborated gDNA results (see FIG. 13D). Finally, a
representative sample of each cellular suspension was treated with red blood
cell lysis buffer
before quantification of cell number with an automated counter and
visualization of cells
under phase contrast microscopy. Cell counts, which primarily reflected single
cells but may
also include some small clusters, were similar to gDNA results (FIG. 16B). The
main
difference was that liver now provided values that were comparable to kidney.
This suggests
that a significant portion of kidney cells may have remained in aggregates
that could have
passed through the cell strainer and been lysed to obtain gDNA. Alternatively,
the cell
counter may have detected more debris in liver suspensions, which was seen in
micrographs
for both minced controls and device treated samples (FIG. 16C).
[0073] Analysis of Cell Types, Numbers, and Viability using Flow Cytometry.
The
final evaluation focused on determining single cell numbers and viability.
Fresh mouse
kidney and liver samples were prepared and digested as described in the
previous section,
except the un-minced control was removed and a 10 min device treatment was
added.
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Digested cellular suspensions were filtered through a 40 p.m cell strainer and
labeled with a
panel of four fluorescent probes: CellMaskTm Green to stain phospholipid cell
membranes,
Draq5 to stain DNA within all cells, 7AAD to stain DNA only within dead cells
with
disrupted plasma membranes, and CD45 to stain leukocytes (Table 1 below).
Table 1
Ce11MaskTm Green Draq5 CD45-PE 7AAD
Assay
(Lipid Membrane) (Nucleus) (Leukocytes) (Dead Cells)
Red blood cells
Leukocytes +/-
Tissue cells +/-
[0074] This panel enabled distinction of tissue cells from non-cellular
debris, anucleated
red blood cells, and leukocytes, while simultaneously assessing viability.
Stained cell
suspensions were analyzed with a BD AccuriTM Flow Cytometer to obtain the
number of each
cell type using the gating protocol described in the methods section and shown
in FIG. 17.
Comparing the relative numbers for each cell type (FIG. 18A and 18B), red
blood cells
constituted the majority of all but the minced control that was digested for
15 min.
Unexpectedly, red blood cell percentage increased slightly as the tissue was
digested more
thoroughly, although this effect was not significant. Leukocyte percentage
remained stable,
decreasing slightly with digestion time. Tissue cell counts, which are
expected to
predominantly be epithelial, were quantified for kidney and liver samples and
are presented
in FIG. 18C and 18D; respectively. Tissue cell numbers were 2 to 5 times
higher for kidney
than liver for the minced controls, which both increased with digestion time.
The increases
were more than an order of magnitude between 15 to 30 min, and 5-fold between
30 to 60
min. With device treatment, there was little change between 10 and 15 min time
points,
although 10 min was associated with high variability for kidney samples.
Extending
processing time to 30 min increased cell number by only ¨50% for both tissue
types,
although differences were not significant. Compared to the minced controls,
device treatment
again provided superior results at the same digestion time point. For kidney,
cell number
differences were 30-fold at 15 min and 4-fold at 30 min. Differences were
about half these
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values for liver. Furthermore, 15 min device treatment yielded similar or
better results than
the minced control that was digested for 30 min. However, the minced control
that was
digested for 60 min now provided the highest cell numbers, exceeding the 30
min device
treatment by 50% for kidney and 100% for liver. This finding is in contrast to
the gDNA
results, particularly for kidney, but generally consistent with CyQUANTO and
cell counter
data. Thus, a significant portion of new cells that are liberated by the
digestion device likely
reside within small aggregates or clusters, which would be reasonable
considering the
smallest channel feature size is 200 um. Finally, viability was assessed using
a DNA dye that
is excluded from healthy cells with intact membranes. Viability was
approximately 80% for
all kidney samples except the minced control that was digested for 60 min and
30 min device
cases, which both dropped to 70% (see FIG. 19). For liver, viability was
approximately 90%
for the minced controls, 80% for 10 and 15 min device treatments, and 70% for
30 min
device treatment. The number of live tissue cells obtained from each condition
is also
presented in FIGS. 18C and 18D. For kidney, 30 min device treatment produced
approximately the same number of live single tissue cells as the minced
control that was
digested for 60 min. The 10 and 15 min device treatments produced around half
of this value,
but in a fraction of the time. For liver, the number of live, single tissue
cells did not increase
with device treatment beyond 10 min. This was likely due to the fragile nature
of liver cells,
which may have been damaged or fully destroyed while recirculating through the
device.
Overall, the microfluidic digestion device performed better for kidney samples
despite the
fact that this tissue type is generally considered to be more difficult to
dissociate. This is
likely due to the combination of kidney cells being more robust and the denser
kidney tissue
requiring higher shear forces to be dissociated.
[0075] A microfluidic device 10 is disclosed that is used to extract or
isolate single cells
from cm x mm-scale tissues using the combination of hydrodynamic shear forces
and
proteolytic digestion. Upon testing of the microfluidic digestion device with
kidney and liver
tissue samples, improvements in recovery of DNA and single tissue cells were
consistently
observed relative to standard methods that require mincing with a scalpel.
Device
performance at short processing times was particularly exciting, as a 10 min
treatment
yielded results that were within 50% of scalpel mincing and digesting for 1
hour, but with
improved viability. Recovery improvements were most striking for DNA,
suggesting that the
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current device design may have left a significant number of cells within small
aggregates or
clusters. Improvements in device function and operation may be found in
improving hydro-
mincing such as decreasing channel dimensions, increasing flow rate, and
installing valves 54
such as illustrated in FIG. 10 to direct flow to different regions of the
tissue. These
approaches could also improve aggregate dissociation of tissue. In addition,
the digestion
device may be paired with another dissociation device 100 such as that
illustrated in FIG. 2
such as the branching channel array with hydrodynamic micro-scalpels. There
was some
observation of evidence that cells may have been damaged during initial tissue
digestion, or
more likely while repeatedly recirculating through the device, particularly
for liver. Thus,
next generation designs will seek to remove single cells as soon as they are
liberated via
filtration or another means of physical separation rather than recirculation.
The microfluidic
device 10 may be used with other tissues such as solid tumors from various
cancer types for
diagnostic purposes and other healthy tissues such as skin, heart, and fat for
use in tissue
engineering and regenerative medicine. That is to say, various tissue types
may be used in
connection with the microfluidic device 10. This includes diseased tissue such
as cancerous
tissue or it may include healthy tissue. Moreover, the tissues or samples may
be obtained
from a number of different organs or tissue types.
[0076] EXAMPLE
[0077] The following is a non-limiting example of the present invention. It
is to be
understood that said example is not intended to limit the present invention in
any way.
Equivalents or substitutes are within the scope of the present invention.
[0078] Fluid Dynamics Simulations. Flow profiles within device channels (as
illustrated
in FIG. 7) were simulated using COMSOL Multiphysics software. This involved
coupling the
Navier-Stokes equations and the continuity equation in finite element fluid
dynamics
simulations. Fluid flow was assumed to be laminar, and the no-slip boundary
condition was
enforced at the channel walls. A flow rate of 1 mL/min was used, but flow
profiles remain the
same at different flow rates up to 20 mL/min as used for experiments. The only
difference is
a corresponding change in maximum flow velocity.
[0079] Device Fabrication. Digestion devices were designed using Onshape
software.
Fluidic channels and hose barb openings were laser etched using a VLS 4.60 60W
CO2 laser
(Universal Laser Systems, Scottsdale, AZ). Channel designs were etched in 6" x
6" optically
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clear cast acrylic sheets (McMaster-Carr, Elmhurst, IL) that served as the
bottom layer of the
device. Hose barb openings were then tapped to provide threading. A gasket was
prepared
from PDMS (Ellsworth Adhesives, Germantown, WI) by casting a 5 mm slab and
cutting
with a scalpel. The device was assembled with the PDMS gasket placed between
the top and
bottom acrylic layers, and secured with nylon screws. The inlet and outlet of
the device were
connected to a peristaltic pump that was controlled by a custom-built Arduino
Uno R3
microcontroller.
[0080] Tissue models. Beef liver was purchased from a local butcher and
tissue cores
were extracted by using a Tru-CutTm biopsy needle (CareFusion, Vernon Hills,
IL) in a
manner analogous to obtaining a clinical biopsy. Briefly, the obturator was
retracted to cover
the specimen notch and the cannula handle was held firmly while the needle was
inserted into
the tissue. The obturator was quickly advanced as far as permitted to position
the specimen
notch in the tissue and the cannula handle was quickly advanced to cut the
tissue. Tissue
obtained in the specimen notch was then transferred to device using tweezers.
Mouse liver
and kidneys were harvested from sacrificed C57B/6 or BALB/c mice (Jackson
Laboratory,
Bar Harbor, ME) that were deemed waste from a research study approved by the
University
of California, Irvine, Institutional Animal Care and Use Committee (courtesy
of Dr. Angela
G. Fleischman). Animal organs were cut with a scalpel into 1 cm long x 1 mm
diameter
pieces, and the mass of each was recorded. Mouse kidneys were sliced in a
symmetrical
fashion to obtain histologically similar portions that included both cortex
and medulla.
[0081] Digestion of tissue samples. The digestion device was first primed
with 200 pL
collagenase type I (Stemcell Technologies, Vancouver, BC) and heated to 37 C
inside an
incubator to ensure optimal enzymatic conditions. Tissue was then placed
inside the chamber
before the device was assembled, secured with nylon screws, and filled with 1
mL
collagenase. Experiments were initiated by flowing fluid through the device at
20 mL/min
with the peristaltic pump, and every 5 min the flow was reversed to clear
tissue from the
downstream sieve gates. Device effluents were collected by pumping directly
into a conical
tube. Controls were digested in a conical tube that contained 1 mL
collagenase, either with or
without prior mincing with a scalpel into ¨1 mm3 pieces. Tubes were placed
inside a 37 C
incubator and gently agitated on a rotating mixer. Every 5 min, the tubes were
vortexed to
mechanically disrupt tissue and maximize digestion. At the conclusion of
digestion
24
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procedures, all cell suspensions were repeatedly vortexed and pipetted to
mechanically
disrupt aggregates and treated with DNase 1(10 pL; Roche, Indianapolis, IN) at
37 C for 5
min.
[0082] Image analysis to monitor tissue digestion. During device operation,
images of
the tissue were captured every 5 min using a camera mounted directly above the
device as
illustrated in FIG. 6. Raw images were processed using ImageJ by first
converting to binary
to identify the borders of the tissue (see FIG. 14). Mean gray value was then
determined
within the tissue border, and multiplied by the area to obtain a single metric
accounting for
tissue size and density. Results at each time point were normalized by the
initial value prior
to the experiment, and presented as percent tissue remaining.
[0083] Quantification of DNA recovered from cell suspensions. DNA content of
digested cell suspensions was assessed by extraction and purification, as well
as direct
assessment within cells using a fluorescent DNA stain. For both cases, samples
were first
filtered using a 70 pm cell strainer to remove remaining tissue and large
aggregates. Purified
genomic DNA (gDNA) was isolated using the QIAamp0 DNA Mini Kit (Qiagen,
Germantown, MD) according to manufacturer's instructions and quantified using
a Nanodrop
ND-1000 (Thermo Fisher, Waltham, MA). DNA within cells was labelled using the
CyQUANTO NF Cell Proliferation Assay Kit (Thermo Fisher, Waltham, MA)
according to
the manufacturer's instructions. Briefly, samples were suspended in HBSS
supplemented
with 35mg/L sodium bicarbonate and 20 mM HEPES and added to an opaque 96-well
plate
(Corning, Corning, NY) in triplicate. An equal volume of CyQUANTO dye was then
added
to each well, incubated at 37 C for 40 minutes under continuous mixing at 200
RPM, and
fluorescence signal was quantified using a Synergy 2 plate reader (BioTek,
Winooski, VT).
Wells containing only HBSS and CyQUANTO dye were used for background
subtraction.
gDNA and fluorescence intensities were normalized by the initial tissue mass.
[0084] Cell counting and imaging of cell suspensions. Digested effluents
were collected,
filtered using a 70 im cell strainer, and incubated with red blood cell lysis
buffer containing
ammonium chloride, potassium carbonate, and EDTA (Biolegend, San Diego, CA)
for 5 min
at room temperature. Cell concentration was determined using a Moxi Z cell
counter with
type S cassettes (Orflo, Hailey, ID), and converted to cell number per mass of
tissue using the
total volume recovered and the initial tissue mass. Imaging was performed by
transferring
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samples to a 12-well plate, waiting for 1 hour for the cells to settle, and
capturing images
using a Hoffman phase contrast microscope with a 4x objective.
[0085] Flow cytometric analysis of single cells. Digested mouse kidney and
liver cell
suspensions were evenly divided into FACS tubes (Corning, Corning, NY) and
resuspended
in FACS Buffer (1X PBS, pH 7.4 without Ca and Mg cations) supplemented with 1%
BSA
and 0.1% NaN3. Samples were first stained with 0.5X CellMaskTm Green (Thermo
Fisher,
Waltham, MA) and 2.5pg/mL anti-mouse CD45-PE monoclonal antibody (clone 30-
F11,
(BioLegend, San Diego, CA) for 20 minutes at 37 C and washed twice with FACS
Buffer by
centrifugation. Cells were then resuspended in FACS buffer supplemented with
12.5 p.M
Draq5 (BioLegend, San Diego, CA) and 5 p.g/mL 7AAD (BD Biosciences, San Jose,
CA)
and maintained on ice for at least 15 minutes prior to analysis on an Accuri
Flow Cytometer
(BD Biosciences). An isotype matched, PE-conjugated monoclonal antibody (clone
RTK4530, BioLegend, San Diego, CA) was used as a control. Flow cytometry data
was
compensated and analyzed using FlowJo software (FlowJo, Ashland, OR).
Compensation
was determined using the kidney and liver tissues that were minced with a
scalpel and
digested for 60 mi, which were aliquoted into four different preparations to
obtain distinct
positive and negative subsets for each probe. The four preparations included
cell fractions
with: 1) negative control CompBeads (3.0-3.4 p.m diameter, BD Biosciences, San
Jose, CA)
and CellMaskTm Green membrane stain, 2) RBCs lysed and CD45-PE antibody, 3)
live and
dead (heat-killed at 55 C for 30 min) cells with 7AAD stain, and 4) Draq5
stain. Gates
encompassing the positive and negative subpopulations within each compensation
sample
were inputted into FlowJo to automatically calculate the compensation matrix.
Finally, a
sequential gating scheme was used to identify different cell subpopulations.
(see FIG. 17). A
SSC-A vs. FSC-A gate was created to select all cellular events and exclude
debris from
further analysis. Multicellular aggregates were removed from the analysis
population to focus
only on single cells using an FSC-H vs. FSC-A gate. Leukocytes were first
distinguished
from the single cell population based on CD45 expression (FL2-A or PE vs. SSC-
H).
Anucleate red blood cells were distinguished by their absence of Draq5 nuclear
stain (FL4-A
or Draq5 vs. SSC-H). The cellularity of the final remaining single cells (CD45
negative,
Draq5 positive) was confirmed by detecting cell membranes using CellMaskTm
Green stain
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(FL1-A or CellMaskTm green vs. FSC-H). Finally, live and dead nucleated tissue
cell
percentages were discriminated based on 7AAD signal (FL3-A or 7AAD vs. SSC-H).
[0086] Statistics. Data are represented as the mean standard error
determined from at
least three independent experiments. P-values were calculated using students t-
test based on
the mean and standard error between different experimental conditions.
[0087] While embodiments of the present invention have been shown and
described,
various modifications may be made without departing from the scope of the
present
invention. For example, aspects of one embodiment may be used in connection
with other
embodiments even though such substitution or combination is not explicitly
described herein.
Further, the publication Qiu et al., Microfluidic device for rapid digestion
of tissues into
cellular suspensions, Lab Chip, 17, 3300 (2017) and its supplementary
information is
incorporated by reference herein. Although there has been shown and described
the preferred
embodiment of the present invention, it will be readily apparent to those
skilled in the art that
modifications may be made thereto which do not exceed the scope of the
appended claims.
Therefore, the scope of the invention is only to be limited by the following
claims. In some
embodiments, the figures presented in this patent application are drawn to
scale, including the
angles, ratios of dimensions, etc. In some embodiments, the figures are
representative only
and the claims are not limited by the dimensions of the figures.
[0088] The reference numbers recited in the below claims are solely for
ease of
examination of this patent application, and are exemplary, and are not
intended in any way to
limit the scope of the claims to the particular features having the
corresponding reference
numbers in the drawings.
[0089] The invention, therefore, should not be limited, except to the
following claims, and
their equivalents.
27
