Patent Application
20240166723
Tumors are often described as “wounds that do not heal” (Dvorak, The New England Journal of Medicine, 1986, 315) since tumorigenesis is tightly associated with chronic tissue damage and repair processes (Arwert et al., Nature Reviews Cancer, 2012, 12). The hallmarks of cancer highly overlap with those of wound healing (MacCarthy-Morrogh, Science Signaling 2020, 13, eaay8690), both featuring sustained proliferative signaling, cell migration, angiogenesis, immune activation and inflammation and tissue repair with extracellular matrix (ECM) remodeling (MacCarthy-Morrogh, Science Signaling, 2020, 13, eaay8690).
In scar tissue formation, after initial blood clots are formed, there is a rapid recruitment of neutrophils and monocytes, with the onset of inflammation. Meanwhile, activation of myofibroblasts induces wounded tissue contraction and increased ECM deposition and remodeling, followed by re-epithelialization and angiogenesis to restore tissue homeostasis (Xue & Jackson, Advances in Wound Care: The Journal For Prevention And Healing, 2015, 4). However, cancer cells harbor high genomic instability and resistance to cell death, which fundamentally differentiate tumor progression from normal wound healing (MacCarthy-Morrogh, Science Signaling, 2020, 13, eaay8690; Behan et al., Nature, 2019, 568). Solid tumors and fibrotic diseases include most of these processes, but in a highly dysregulated and tissue dependent manner. Importantly, pathological ECM remodeling (Cox & Erler, Disease Models & Mechanisms, 2011, 4; Yuzhalin et al., Biochimica Et Biophysica Acta, Reviews On Cancer, 2018, 1870) is usually implicated in liver carcinoma (Nguyen et al., Communications Biology, 2022, 5, 202), breast cancer (Conklin et al., The American Journal Of Pathology, 2011, 178, 1221), melanoma (Kaur et al., Cancer Discovery, 2019, 9, 64) and lung cancer (Wood et al., Cancer Treatment Reviews, 2014, 40, 558). The dysregulated ECM signatures are directly associated with poor prognosis and immunotherapy failure (Chakravarthy et al., Nature Communications, 2018, 9, 4692).
3D in vitro models have been used to mimic specific aspects of the tumor microenvironment (TME) and study the dynamic ECM evolution (Pizzurro et al., Cancers, 2021, 13). Previous work on reconstructing the tumor ECM in vitro has primarily focused on individual local parameters, such as pore size, fiber diameter (Seo et al., Proceedings Of The National Academy Of Sciences Of The United States Of America, 2020, 117, 11387; Kalbitzer & Pompe, Acta Biomaterialia, 2018, 67, 206; Sapudom et al., Biomaterials, 2015, 52, 367; Yang et al., Biomaterials, 2010, 31, 5678), stiffness (Mason et al., Acta Biomaterialia, 2013, 9, 4635), viscoelasticity (Wisdom et al., Nature Communications, 2018, 9, 4144), collagen alignment (Riching et al., Biophysical Journal, 2014, 107, 2546; Mosier et al., Biophysical Journal, 2019, 117, 1692) or microarchitecture (Fischer et al., Proceedings Of The National Academy Of Sciences Of The United States Of America, 2021, 118; Sakar et al., Nature Communications, 2016, 7, 11036; Guzman et al., Biomaterials 2014, 35, 6954). However, these models usually only include sub-micrometer topographies and mechanics but lack global architectural similitude to in vivo ECM structure.
Thus, there is a need in the art for a novel method of generating thickened collagen bundles and clusters. The present invention satisfies this unmet need.
In one aspect, the present invention relates to a method of generating thickened collagen bundles or clusters, comprising the steps of: preparing a neutralized collagen gel sample; warming the neutralized collagen gel sample to provide a warmed neutralized collagen gel sample; and agitating the warmed neutralized collagen gel sample to provide an agitated collagen gel sample.
In one embodiment, the method further comprises the steps of: cooling the agitated collagen gel sample to about 0° C. for about 10 to 30 minutes to provide a cooled collagen gel sample; and reagitating the cooled collagen gel sample at a temperature greater than or equal to 0° C. In one embodiment, the method further comprises the step of allowing the neutralized collagen gel sample to sit at room temperature for 6 to 10 minutes. In one embodiment, the method further comprises the step of cooling the neutralized collagen gel sample to 0° C.
In one embodiment, the step of warming the neutralized collagen gel sample comprises the step of heating the neutralized collagen gel sample to a temperature greater than or equal to 20° C. In one embodiment, the step of agitating the warmed neutralized collagen gel sample comprises the step of mixing the warmed neutralized collagen gel sample with a pipette.
In one embodiment, the agitated collagen gel sample has a turbidity which measures above background. In one embodiment, the step of agitating the warmed neutralized gel sample is automated via a mixing device.
In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 10 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel provides disconnected collagen clusters.
In one embodiment, the method further comprises the step of mixing the disconnected collagen clusters with cells to provide a cell and collagen cluster mixture. In one embodiment, the cell and collagen cluster mixture self-aggregates. In one embodiment, the cell and collagen cluster mixture comprises a structure selected from the group consisting of compact cell-collagen clusters, spheroids, and fibrotic cell clusters.
In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 10 mg/mL.
In one embodiment, the neutralized collagen sample comprises a collagen selected from the group consisting of type I collagen, type II collagen, type III collagen, type IV collagen, type XI collagen, thick collagen, hydrolyzed collagen, mammal collagen, powdered collagen, and synthetic collagen. In one aspect, the present invention provides a thickened collagen structure.
In one embodiment, the thickened collagen structure is macroporous or mesoporous. In one embodiment, the thickened collagen structure is used in a bioactive cue, an in situ injectable, a vaccine, a biomimetic material, or a vascular organoid. In one embodiment, the thickened collagen structure comprises at least one collagen island. In one embodiment, the area of the at least one collagen island is between 1 and 1500 μm2.
The foregoing purposes and features, as well as other purposes and features, will become apparent with reference to the description and accompanying figures below, which are included to provide an understanding of the invention and constitute a part of the specification, in which like numerals represent like elements.
It is to be understood that the figures and descriptions of the present invention have been simplified to illustrate elements that are relevant for a clear understanding of the present invention, while eliminating, for the purpose of clarity, many other elements found in related systems and methods. Those of ordinary skill in the art may recognize that other elements and/or steps are desirable and/or required in implementing the present invention. However, because such elements and steps are well known in the art, and because they do not facilitate a better understanding of the present invention, a discussion of such elements and steps is not provided herein. The disclosure herein is directed to all such variations and modifications to such elements and methods known to those skilled in the art.
Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention belongs. Although any methods and materials similar or equivalent to those described herein can be used in the practice or testing of the present invention, exemplary methods and materials are described.
As used herein, each of the following terms has the meaning associated with it in this section.
The articles “a” and “an” are used herein to refer to one or to more than one (i.e., to at least one) of the grammatical object of the article. By way of example, “an element” means one element or more than one element.
“About” as used herein when referring to a measurable value such as an amount, a temporal duration, and the like, is meant to encompass variations of ±20%, ±10%, ±5%, ±1%, and ±0.1% from the specified value, as such variations are appropriate.
Throughout this disclosure, various aspects of the invention can be presented in a range format. It should be understood that the description in range format is merely for convenience and brevity and should not be construed as an inflexible limitation on the scope of the invention. Accordingly, the description of a range should be considered to have specifically disclosed all the possible subranges as well as individual numerical values within that range. For example, description of a range such as from 1 to 6 should be considered to have specifically disclosed subranges such as from 1 to 3, from 1 to 4, from 1 to 5, from 2 to 4, from 2 to 6, from 3 to 6 etc., as well as individual numbers within that range, for example, 1, 2, 2.7, 3, 4, 5, 5.3, 6 and any whole and partial increments therebetween. This applies regardless of the breadth of the range.
The term “extracellular matrix” as used herein refers to a scaffold in a cell's external environment with which the cell interacts via specific cell surface receptors. The extracellular matrix serves many functions, including, but not limited to, providing support and anchorage for cells, segregating one tissue from another tissue, and regulating intracellular communication. The extracellular matrix is composed of an interlocking mesh of fibrous proteins and glycosaminoglycans (GAGs). Examples of fibrous proteins found in the extracellular matrix include collagen, elastin, fribronectin, and laminin. Examples of GAGs found in the extracellular matrix include proteoglycans (e.g., heparin sulfate), chondroitin sulfate, keratin sulfate, and non-proteoglycan polysaccharide (e.g., hyaluronic acid).
As used herein, the term “spheroid” refers to microtissues of cells growing and/or interacting within their surroundings in all three dimensions in an artificially-created environment. Such microtissues can comprise a plurality of homotypic or heterotypic cells, preferably mammalian cells, more preferably human cells. Such 3-D cell cultures more closely resemble the in vivo surroundings of the cells as compared to 2-D cell cultures. Spheroids provide a more accurate model system for cellular, physiological and/or pharmaceutical studies than cells grown in conventional two-dimensional cultures.
As used herein, the term “fibrotic cell” is defined as an injured cell that has a lower capability to conduct or generate an electrical impulse as compared to normal, healthy cells. Such fibrotic cell has lost its capability to conduct or generate an electrical impulse. Fibrotic cells are obtained in various ways. Surrounding cells can be rendered fibrotic by heating them with a heating element with a temperature of at least 55° C., or cooling them with a cooling element with a temperature of at most −75° C.
“Collagen” is a structural protein found in connective tissue; it frequently takes the form of fibrils arranged in a triple helix. Fibrillar types of collagen include Types I, II, III, V, and XI. Type I collagen makes up a great deal of the organic part of bone as well as being found in skin, tendons, blood vessels, and organs, while type III collagen is commonly found near or with type I. On the other hand, cartilage is composed primarily of type II collagen. Other types of cartilage are less common and may be found in membranes, on cell surfaces, and associated with hair and placental structures.
In one aspect, the present invention relates to a method of generating thickened collagen bundles, the method comprising the steps of preparing a neutralized collagen gel sample; warming the gel sample; and agitating the warmed gel sample until the gel sample has a turbidity above background. In one embodiment, the method generates clusters. Any embodiment involving a bundle is applicable to any embodiment involving a cluster, and vice-versa.
In one embodiment, the method may be employed to prepare collagen through the modulation of collagen-based ECM scaffolds by disrupting the gelation process. In one embodiment, the method is performed manually. In one embodiment, the method is performed using an automated device. In one embodiment, the method is performed using a mixing device.
In one embodiment, the collagen is derived from rat tail. In one embodiment, the collagen is derived from bovine. In one embodiment, the collagen has been neutralized.
In one embodiment, the method involves cooling the collagen gel to about 0° C. In one embodiment, the method involves cooling the collagen gel to about 0° C. to 5° C. In one embodiment, the method involves cooling the collagen gel to about 5° C. to 10° C. In one embodiment, the method involves cooling the collagen gel to about 10° C. to 15° C. In one embodiment, the method involves cooling the collagen gel to about 15° C. to 20° C. In one embodiment, the method involves cooling the collagen gel to about 20° C. to 25° C.
In one embodiment, the method involves warming the cooled collagen gel to about 0° C. In one embodiment, the method involves warming the cooled collagen gel to about 5° C. In one embodiment, the method involves warming the cooled collagen gel to about 10° C. In one embodiment, the method involves warming the cooled collagen gel to about 15° C. In one embodiment, the method involves warming the cooled collagen gel to about 20° C. In one embodiment, the method involves warming the cooled collagen gel to about 25° C. In one embodiment, the method involves warming the cooled collagen gel to ambient temperature.
In one embodiment, the method involves letting the warmed collagen gel sit for 6 to 7 minutes. In one embodiment, the method involves letting the warmed collagen gel sit for 6 to 8 minutes. In one embodiment, the method involves letting the warmed collagen gel sit for 6 to 9 minutes. In one embodiment, the method involves letting the warmed collagen gel sit for 6 to 10 minutes. In one embodiment, the method involves letting the warmed collagen gel sit for 5 to 10 minutes. In one embodiment, the method involves letting the warmed collagen gel sit for 5 to 8 minutes. In one embodiment, the method involves letting the warmed collagen gel sit for 5 to 9 minutes.
In one embodiment, the method involves warming the gel after sitting to about 25° C. In one embodiment, the method involves warming the gel after sitting to about 30° C. In one embodiment, the method involves warming the gel after sitting to about 31° C. In one embodiment, the method involves warming the gel after sitting to about 32° C. In one embodiment, the method involves warming the gel after sitting to about 33° C. In one embodiment, the method involves warming the gel after sitting to about 34° C. In one embodiment, the method involves warming the gel after sitting to about 35° C. In one embodiment, the method involves warming the gel after sitting using heat from a hand.
In one embodiment, the method involves pipetting the warmed gel. In one embodiment, the method involves mixing the warmed gel with a pipette. In one embodiment, the method involves mixing the warmed gel with an automated device. In several embodiments, mixing the warmed gel increases turbidity. In one embodiment, the warmed gel is mixed until it has a turbidity above background.
In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 5 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 10 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 15 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 20 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 30 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 40 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 50 minutes. In one embodiment, the step of agitating the warmed neutralized collagen gel is performed for over about 60 minutes.
In one embodiment, the method provides disconnected collage clusters. In one embodiment, the method provides loosely connected collagen clusters. In one embodiment, the method further comprises the step of mixing collagen clusters with cells. In one embodiment, the method provides a cell and collagen cluster mixture.
In one embodiment, the method further comprises the step of placing the cell and collagen cluster mixture onto a low adhesion plate. In one embodiment, the method further comprises the step of placing the cell and collagen cluster mixture into a well. In one embodiment, the method further comprises the step of forming the cell and collagen cluster mixture into hanging droplets. In one embodiment, the step of placing the cell and collagen cluster mixture is performed with a pipette. In one embodiment, the step of forming the cell and collagen cluster mixture is performed with a pipette.
In one embodiment, the cell and collagen cluster mixture self-aggregates. In one embodiment, the cell and collagen cluster mixture undergo cell aggregation. In one embodiment, the cell and collagen cluster mixture comprises a structure selected from the group consisting of compact cell-collagen clusters, spheroids, and fibrotic cell clusters.
In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 10 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 9 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 8 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 7 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 6 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 5 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 4 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 3 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 1 mg/mL to about 2 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 10 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 9 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 8 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 7 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 6 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 5 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 4 mg/mL. In one embodiment, the neutralized collagen sample has a collagen concentration of about 2 mg/mL to about 3 mg/mL.
In one embodiment, the neutralized collagen gel sample comprises a collagen selected from the group consisting of type I collagen, type II collagen, type III collagen, type IV collagen, type XI collagen, thick collagen, hydrolyzed collagen, mammal collagen, powdered collagen, and synthetic collagen.
In one embodiment, turbidity of the gel is measured with a microplate reader. In one embodiment, the microplate reader is pre-heated. In one embodiment, the colorimetric readout is monitored. In one embodiment, the turbidity is determined by the colorimetric readout.
In one embodiment, the microplate reader is pre-heated to 30° C. In one embodiment, the microplate reader is pre-heated to 31° C. In one embodiment, the microplate reader is pre-heated to 32° C. In one embodiment, the microplate reader is pre-heated to 33° C. In one embodiment, the microplate reader is pre-heated to 34° C. In one embodiment, the microplate reader is pre-heated to 35° C. In one embodiment, the microplate reader is pre-heated to 36° C. In one embodiment, the microplate reader is pre-heated to 37° C. In one embodiment, the microplate reader is pre-heated to 38° C. In one embodiment, the microplate reader is pre-heated to 39° C. In one embodiment, the microplate reader is pre-heated to 40° C.
In one embodiment, the colorimetric readout is recorded every 1 minute for 20 minutes. In one embodiment, the colorimetric readout is recorded every 1 minute for 25 minutes. In one embodiment, the colorimetric readout is recorded every 1 minute for 30 minutes. In one embodiment, the colorimetric readout is recorded every 1 minute for 35 minutes. In one embodiment, the colorimetric readout is recorded every 1 minute for 40 minutes. In one embodiment, the colorimetric readout is recorded every 2 minutes for 20 minutes. In one embodiment, the colorimetric readout is recorded every 5 minutes for 20 minutes.
In one embodiment, the stirring of the warmed gel results in a turbidity around 0.4-0.5. In one embodiment, the stirring of the warmed gel results in a turbidity around 0.4-0.6. In one embodiment, the stirring of the warmed gel results in a turbidity around 0.4-0.7. In one embodiment, the stirring of the warmed gel results in a turbidity around 0.4-0.8. In one embodiment, the stirring of the warmed gel results in a turbidity around 0.4-0.9. In one embodiment, the stirring of the warmed gel results in a turbidity around 0.4-1.0.
In another aspect, the present invention provides thick collagen bundle gels. In one embodiment, the thick collagen bundle gels are globally softer than normal collagen gels. In one embodiment, the thick collagen bundle gels display viscoelastic behavior similar to that of normal collagen gels. In one embodiment, the thick collagen bundle gels display higher frequency dependent behavior than normal collagen gels during bulk rheometry.
In one embodiment, the thick collagen bundle gels possess stiff mesoscopic clusters. In one embodiment, the thick collagen bundle gels have tunable mesoscopic patterns. In one embodiment, the thick collagen bundle gels contain pores. In one embodiment, the thick collagen bundle gels display macroporosity. In one embodiment, the thick collagen bundle gels display mesoporosity.
In one embodiment, the diameters of the pores range from 1 to 10 μm. In one embodiment, the diameters of the pores range from 1 to 20 μm. In one embodiment, the diameters of the pores range from 1 to 30 μm. In one embodiment, the diameters of the pores range from 1 to 40 μm. In one embodiment, the diameters of the pores range from 1 to 50 μm. In one embodiment, the diameters of the pores range from 10 to 20 μm. In one embodiment, the diameters of the pores range from 10 to 30 μm. In one embodiment, the diameters of the pores range from 10 to 40 μm. In one embodiment, the diameters of the pores range from 10 to 50 μm. In one embodiment, the diameters of the pores range from 10 to 60 μm. In one embodiment, the diameters of the pores range from 10 to 70 μm. In one embodiment, the diameters of the pores range from 10 to 80 μm. In one embodiment, the diameters of the pores range from 10 to 90 μm. In one embodiment, the diameters of the pores range from 10 to 100 μm. In one embodiment, the diameters of the pores range from 100 to 200 μm. In one embodiment, the diameters of the pores range from 200 to 300 μm. In one embodiment, the diameters of the pores range from 300 to 400 μm. In one embodiment, the diameters of the pores range from 400 to 500 μm.
In one embodiment, the thick collagen bundle gels can be incorporated with several cell types. Exemplary cell types include, but are not limited to, stem cells and tissue cells to study development. In one embodiment, cancer cells are embedded into the hydrogels to study dissemination in physiologically relevant settings. In one embodiment, cancer cells are embedded into the hydrogels to study migration dynamics in physiologically relevant settings. In one embodiment, the hydrogels are applied to study differentiation in vitro. In one embodiment, the thick collagen bundle gels can be incorporated with co-gel biomaterials.
In one embodiment, the thick collagen bundle gels are used in three-dimensional tissues. In one embodiment, the thick collagen bundle gels are used in a vascular organoid.
In one embodiment, the thick collagen bundle gels are used to induce stem cell differentiation. In one embodiment, the thick collagen bundle gels are used to induce mesodermal differentiation.
In one embodiment, the thick collagen bundle gels can serve as bioactive cues to influence the behavior of cells. In one embodiment, the thick collagen bundle gels facilitate cell infiltration. In one embodiment, the thick collagen bundle gels are employed in in situ injectables. In one embodiment, the thick collagen bundle gels are employed in vaccines. In one embodiment, the thick collagen bundle gels are used as microporous gels.
In one embodiment, the thick collagen bundle gels are used for immune stimulation. In one embodiment, the thick collagen bundle gels are used for wound healing. In one embodiment, the thick collagen bundle gels are used for vascularization.
In one embodiment, the hydrogels contain mesoscaled architectures. In one embodiment, the hydrogels contain macroscaled architectures. In one embodiment, the hydrogels are clusters.
In another aspect, the present invention provides collagen island hydrogels. In one embodiment, the collagen island hydrogels require only mechanical force to form. In one embodiment, the collagen island hydrogels are formed using shear. In one embodiment, the collagen island hydrogels have tunable island size. In one embodiment, the island size is dependent on mechanical shear frequency. In one embodiment, the collagen island hydrogels can be isolated from solution and resuspended in other natural ECM-derived matrices. In one embodiment, the collagen island hydrogels have tunable topography. In one embodiment, the collagen island hydrogels have tunable mechanical properties.
In one embodiment, the collagen island hydrogels are formed via mixing. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 10 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 10 seconds. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 20 seconds. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 30 seconds. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 40 seconds. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 50 seconds. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 60 seconds. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 2 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 3 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 4 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 5 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 6 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 7 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 8 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 second to about 9 minutes. In one embodiment, the collagen island hydrogels are formed via mixing for about 1 minute to about 10 minutes.
In one embodiment, the area of the collagen islands ranges from 1 to 1500 μm2. In one embodiment, the area of the collagen islands ranges from 1 to 1000 μm2. In one embodiment, the area of the collagen islands ranges from 1 to 750 m2. In one embodiment, the area of the collagen islands ranges from 1 to 500 m2. In one embodiment, the area of the collagen islands ranges from 1 to 1000 μm 2 . In one embodiment, the area of the collagen islands ranges from 1 to 450 μm2. In one embodiment, the area of the collagen islands ranges from 1 to 400 μm2. In one embodiment, the area of the collagen islands ranges from 1 to 350 m2. In one embodiment, the area of the collagen islands ranges from 1 to 300 m2. In one embodiment, the area of the collagen islands ranges from 1 to 250 m2. In one embodiment, the area of the collagen islands ranges from 1 to 200 μm2. In one embodiment, the area of the collagen islands ranges from 1 to 150 m2. In one embodiment, the area of the collagen islands ranges from 1 to 100 μm2. In one embodiment, the area of the collagen islands ranges from 1 to 50 μm2.
In several embodiments, the collagen island hydrogels mimic the heterogeneous mesoscopic features seen in tissue. In one embodiment, the collagen island hydrogels are compatible with modular assembly for tissue engineering. In one embodiment, the collagen island hydrogels act as bioactive cues. In one embodiment, the collagen island hydrogels modulate stem cell behavior. In one embodiment, the collagen island hydrogels can promote MSC osteogenesis. In one embodiment, the collagen island hydrogels can direct spontaneous iPSC differentiation. In one embodiment, the collagen island hydrogels can induce cellular self-assembly processes. In one embodiment, the collagen island hydrogels induce microtissue fabrication. In one embodiment, the collagen island hydrogels induce tissue morphogenesis. In one embodiment, the collagen island hydrogels direct cell differentiation. In one embodiment, the collagen island hydrogels direct physical cues in regenerative applications.
In one embodiment, the collagen island hydrogels are used in a biomimetic material. In one embodiment, the biomimetic material displays strain stiffening. In one embodiment, the biomimetic material displays stress relaxation.
In one embodiment, the collagen island hydrogels can be incorporated with several cell types. Exemplary cell types include, but are not limited to, stem cells and tissue cells to study development. In one embodiment, cancer cells are embedded into the hydrogels to study dissemination in physiologically relevant settings. In one embodiment, cancer cells are embedded into the hydrogels to study migration dynamics in physiologically relevant settings. In one embodiment, the hydrogels are applied to study differentiation in vitro. In one embodiment, the collagen island hydrogels can be incorporated with co-gel biomaterials.
In one embodiment, the collagen island hydrogels are used in three-dimensional tissues. In one embodiment, the collagen island hydrogels are used in a vascular organoid.
In one embodiment, the collagen island hydrogels are used to induce stem cell differentiation. In one embodiment, the collagen island hydrogels are used to induce mesodermal differentiation.
In one embodiment, the collagen island hydrogels can serve as bioactive cues to influence the behavior of cells. In one embodiment, the collagen island hydrogels facilitate cell infiltration. In one embodiment, the collagen island hydrogels are employed in in situ injectables. In one embodiment, the collagen island hydrogels are employed in vaccines. In one embodiment, the collagen island hydrogels are used as microporous gels.
In one embodiment, the collagen island hydrogels are used for immune stimulation. In one embodiment, the collagen island hydrogels are used for wound healing. In one embodiment, the collagen island hydrogels are used for vascularization.
In one embodiment, the hydrogels contain mesoscaled architectures. In one embodiment, the hydrogels contain macroscaled architectures. In one embodiment, the hydrogels are clusters. In one embodiment, the hydrogels are thick tortuous fiber bundles.
The invention is further described in detail by reference to the following experimental examples. These examples are provided for purposes of illustration only and are not intended to be limiting unless otherwise specified. Thus, the invention should in no way be construed as being limited to the following examples, but rather, should be construed to encompass any and all variations which become evident as a result of the teaching provided herein.
Without further description, it is believed that one of ordinary skill in the art can, using the preceding description and the following illustrative examples, make and utilize the compositions of the present invention and practice the claimed methods. The following working examples, therefore, specifically point out various embodiments of the present invention, and are not to be construed as limiting in any way the remainder of the disclosure.
Thickened collagen bundles were long and curly in appearance, when compared with short, dense collagen networks gels prepared without mechanical disturbance, i.e. normal collagen, as shown in
Previous studies have focused on the dynamics of fibrillar gel formation. The steps proposed for in vitro fibrillogenesis, portrayed in
As shown in
MDA-MB-231 is a breast cancer triple-negative basal type cell line. It is highly aggressive and metastatic, featuring low-claudin (reduced cell-cell contact) and overexpression of ECM components and remodeling-related proteases, mesenchymal associated markers, and tumor favoring signals (Dai et al., Journal of Cancer 2017, 8, 3131-3141). In in vitro 3D models, MDA-MB-231 cells are highly sensitive to collagen alignment (Nuhn et al., Acta Biomaterialia, 2018, 66, 248-257; Han et al., Proceedings Of The National Academy Of Sciences Of The United States Of America, 2016, 113, 11208-11213), stiffness (Geiger et al., PloS One, 2019, 14, e0225215; Sapudom et al., Biomaterials, 2015, 52, 367-375) and steric hindrance, such as pore size (Wolf et al., The Journal Of Cell Biology, 2013, 201, 1069-1084). In vivo, MDA-MB-231 tumors preferentially metastasize to the bones, brain and lungs (Bos et al., Nature, 2009, 459, 1005-1009; Kang et al., Cancer Cell, 2003, 3, 537-549; Minn et al., The Journal of Clinical Investigation, 2005, 115, 44-55).
The thick collagen system was characterized by reduced physical hindrance, global softness, and local thickened fiber bundles. When embedded in the thick collagen gels, MDA-MB-231 cells responded rapidly during the first 24 h. In long term culture (up to day 5), MDA-MB-231 cells in normal collagen mostly grew into spherical cell clusters whereas the ones seeded in thick collagen remained isolated and rarely clustered (
Persistent cell migration in a 3D environment requires protrusive structures and actomyosin-mediated contractility (Pandya et al., Current Opinion in Cell Biology, 2017, 48, 87-96). The branching and elongation of protrusive structures often involves ARP2/3 and formin-nucleated actin polymerization (Pollard, Annual Review of Biophysics And Biomolecular Structure, 2007, 36, 451-477; Goode & Eck, Annual Review Of Biochemistry, 2007, 76, 593-627), whereas Rho-ROCK signaling is in the center of actomyosin-mediated cell contractility (Amano et al., Cytoskeleton, 2010, 67, 545-554). The Rho GTPases Rho, Rac and Cdc42 are major players in cytoskeletal regulation (Sadok & Marshall, Small GTPases, 2014, 5, e29710). In 3D ECMs, matrix metalloproteinases (MMPs) contribute to matrix degradation and remodeling, which allows cell migration (Singh et al., Frontiers in Molecular Biosciences, 2015, 2, 19; Wolf et al., The Journal Of Cell Biology, 2013, 201, 1069-1084).
To investigate the specific molecular mechanism behind the elevated migration of MDA-MB-231 cells in thick collagen, the following drugs were tested in these gels: pan-MMP inhibitor GM6001 (20 μM), potent selective Cdc42 inhibitor ML141 (10 μM), ROCK inhibitor Y27632 (30 μM), ARP2/3 inhibitor CK666 (50 μM), formin inhibitor SMIFH2 (50 μM) and Rac1 inhibitors NSC23766 (100 μM) and EHT1864 (20 μM). First cell morphology was traced and quantified. GM6001, ML141 and EHT1864 treatment resulted in similar morphology patterns compared with DMSO control in normal collagen. In thick collagen, GM6001, ML141 and EHT1864 treatment also showed similar morphology but still different from the morphology observed in normal collagen conditions (
Notably, each of the observed patterns is directly linked with the cell function directly affected by the inhibitor. Y27632 is a potent and selective ROCK inhibitor. ROCK mediates actomyosin-based cell contractility via regulating retrograde flow of actin monomers from protruding structures (Vicente-Manzanares et al., Nature Reviews. Molecular Cell Biology 2009, 10, 778-790). Inhibition of ROCK in these cultures led to extremely long cell protrusions, as shown in
Nucleation inhibitors triggered distinct morphologies in these collagen cultures. CK666 inhibits function of ARP2/3 whereas SMIFH2 inhibits formins. ARP2/3 often initializes actin fiber branching and is closely associated with lamellipodia. Upon ARP2/3 inhibition, MDA-MB-231 cells, in both normal collagen and thick collagen, were elongated and stretched. This suggested that, in the absence of ARP2/3, formins-driven actin networks may be disrupted and may lead to the elongation of the cell body. However, SMIFH2 treated cells were round and lacked cell peripheral fluctuations (indicated by red arrows on
Further quantification of MDA-MB-231 cells morphology agreed with representative images (
Apart from morphology signatures, cell migration was also tracked (
To conclude, these drug assays together indicate that for highly metastatic MDA-MB-231 cells to migrate in thick collagen, ROCK mediated cell contractility is in the center of tumor cell dissemination. In addition, the role of formins based on SMIFH2 needs to be explored further, since SMIFH2 has been recently reported to have off-target effects on several other non-muscle myosins (Nishimura et al., Journal of Cell Science, 2021, 134). In contrast to the MDA-MB-231 cell line, MCF10A is an immortalized noncancerous breast epithelial cell line. MCF10As lack the potential to form tumors and metastasize in nude mice (Cowell et al., Cancer Genetics and Cytogenetics, 2005, 163, 23-29). When cultured in 3D reconstituted basement membrane, MCF10As undergo differentiation and growth arrest and develop into acinar structures that recapitulate many features of normal breast epithelial cells.
Thick collagen induced different responses from MCF10A single cells compared with both 1 mg/ml or 2 mg/ml normal collagen, but not as dramatic as seen in MDA-MB-231 cells (
Thick collagen also triggered morphological changes of MCF10A acini. Protrusive branches appeared from the acinus and pulled the collagen patches nearby substantially (
The collagen-pulling was not limited to MCF10A acini but also observed in single cells. In overview tile scan imaging (
To investigate the molecular signatures induced by thick collagen in MCF10A cells, immunofluorescence imaging of YAP/TAZ, vinculin, and phosphorylated myosin light chain (pMLC) was collected. YAP/TAZ is a sensitive mechanosensor, and its relative nuclear/cytoplasmic distribution is regulated by substrate rigidity (Dupont et al., Nature, 2011, 474, 179-183). YAP/TAZ accumulates in the nucleus when a cell stretches out on a substrate of high rigidity but localizes in the cytoplasm when cells are in confinement or interacting with softer substrates. As MCF10A cells often clustered in collagen after three days after gel embedding, MCF10A clusters were separated from MCF10A single cells in YAP/TAZ measurement. YAP/TAZ quantification revealed that both MCF10A single cells and MCF10A clusters had a higher nucleus/cytoplasm ratio in thick collagen versus normal collagen (
Vinculin is a molecular marker for both intercellular and cell-ECM adhesions (Bays & DeMali, Cellular and Molecular Life Sciences: CMLS, 2017, 74, 2999-3009). In the protrusive strands from MCF10A cell clusters, vinculin (
pMLC is a direct marker for actomyosin based contractility. Data in both
In this work a fast and reproducible method to modulate collagen architecture was introduced and created a type of collagen scaffold highly similar to in vivo stromal architecture heterogeneity (
There have been many previously reported different ways of modifying collagen architecture for 3D cultures. However, compared with previous work, these collagen scaffolds demonstrated better global resemblance to in vivo ECM architecture (
The instantly presented thick collagen may serve as a better platform to examine and compare cell migration behaviors in 3D, as this type of collagen architecture has decoupled the confounding effect of physical restraints from other factors. Physical limits or steric restraints of collagen greatly impact cell behavior. Most previous reports of tumor cell migration in 3D collagen are based on the normal collagen scaffold, which is strongly impacted by matrix proteolysis (Zaman et al., PNAS 2006, 103, 10889-10894; Sabeh et al., The Journal of Cell Biology, 2004, 167, 769-781). These studies suggest the potential of MMP inhibition as a strategy against invasion. The instantly presented thick collagen suggests an adjusted view of MMP inhibitors that they may be ineffective in heterogeneous and clustered collagen environments (similar to the thick collagen gels presented herein). In addition, the results agree with intravital imaging work (Gligorijevic et al., PLOS Biology, 2014, 12, e1001995) that the migration of breast tumor cells in vivo is MMP-independent and may provide additional indications for the reasons of the failure of MMP inhibitors in clinical trials (Fingleton, The Cancer Degradome, 759-785). Without physical restraints, thick collagen can help further stratify drug performance in in vivo-like microenvironments. For example, in normal collagen, GM6001/ML141/SMIFH2/NSC23766/EHT1864 appear to exert the same effects on cell morphology (all cells are round). However, MDA-MB- 231s in thick collagen fall into two subgroups: GM6001/ML141/EHT1864 versus SMIFH2/NSC23766 (
Thick collagen may serve as a better platform to examine and compare cell migration behaviors in 3D, as this type of collagen architecture has decoupled the confounding effect of physical restraints from other factors. Physical limits or steric restraints (Mosier et al., Biophysical Journal, 2019, 117, 1692-1701); Guzman et al., Biomaterials, 2014, 35, 6954-6963) of collagen greatly impact cell behavior. Most previous reports of tumor cell migration in 3D collagen were based on the normal collagen scaffold, which was strongly impacted by matrix proteolysis (Wolf et al., The Journal Of Cell Biology, 2013, 201, 1069-1084; Zaman et al., Proceedings Of The National Academy Of Sciences Of The United States Of America, 2006, 103, 10889-10894; Sabeh et al., The Journal Of Cell Biology, 2004, 167, 769-781), suggesting MMP inhibition as a strategy against invasion. However, thick collagen suggested an adjusted view of MMP inhibitors since they may be ineffective in heterogeneous and clustered collagen environments such as thick collagen gels. In addition, the results agreed with intravital imaging work (Gligorijevic et al., PLoS Biology, 2014, 12, e1001995) that the migration of breast tumor cells in vivo is MMP-independent and may provide additional indications for the reasons for the failure of MMP inhibitors in clinical trials (in The Cancer Degradome 759-785). Without physical restraints, thick collagen could help further stratify drug performance in in vivo-like microenvironments. For example, in normal collagen, GM6001/ML141/SMIFH2/NSC23766/EHT1864 appeared to exert the same effects on cell morphology. However, MDA-MB-231 cells in thick collagen fell into two subgroups: GM6001/ML141/EHT1864 response versus SMIFH2/NSC23766 response (
Further, this type of collagen can be easily integrated into coculture assays as with normal collagen. The unique architecture and mechanical features of this thick collagen introduce a different overall picture of the mechanical landscape that scaffolds cells in culture. This method captures the spatial heterogeneity in the tumor microenvironment. These findings echoed one intravital imaging study of breast cancer (Gligorijevic et al., PLoS Biology, 2014, 12, e1001995) which identified two subpopulations of tumor cell phenotypes, and their multiparameter classification links the driving factor of these two subpopulations to their relative locations in the TME. This study also supported the importance of spatial heterogeneity of TME. The overall features of this thick collagen system are summarized in
As thick collagen gels were locally dense but globally soft, they provided a way to separate local scale mechanics from global scale (or mesoscale) mechanics. Mechanical testing implied that the thick collagen behaves as a more fluid-like material. Additionally, previous research has shown that stiffness of collagen increases over collagen concentration (Motte & Kaufman, Biopolymers, 2013, 99, 35-46). From the confocal imaging, thick collagen bundles are of higher density compared with thin collagen networks in between. Taken together, it may be possible to conclude that thick collagen networks have a weaker, more fluid-like background but contain stiff inclusions. Previous studies have shown the importance of scales in mechanical interactions. For example, long-distance matrix-mediated mechanical communication exists (Sunyer et al., Science, 2016, 353, 1157-1161; van Oers et al., PLoS Computational Biology 2014, 10, e1003774; Wang et al., Biophysical Journal 2014, 107, 2592-2603; Pakshir et al., Nature Communications, 2019, 10, 1850; Reinhart-King et al., Biophysical Journal 2008, 95, 6044-6051)), fibroblasts can recruit macrophages from 100-200 micrometers away in fibrillar collagen (Wozniak et al., The Journal Of Cell Biology, 2003, 163, 583-595), floating gel regulates differently compared with anchored gel (Grinnell et al., The Journal Of Biological Chemistry 1999, 274, 918-923; Leong et al., Biochemical And Biophysical Research Communications 2010, 401, 287-292)), with the same surface coating, the thickness of the coating regulates cell behavior, tissue-scale deformations can control wound closure (Sakar et al., Nature Communications, 2016), among others. The creation of mesoscale clustered collagen architectures may provide a solution to these problems. Outside tumor modeling, the thick collagen gels have potential applications in 3D wound modeling, stem cell differentiation, and primary cell cultures.
In physiological tissues, ECM topologies and architectures can be diverse, with regions of varying density and stiffnesses (
Fluid dynamics simulations revealed how much shear stress and energy are introduced into gels during this process (
In accordance with simulations, it was found that island size could be reproducibly modulated with 2 second gels having smaller islands and 10 second having larger islands (
These island architectures and their packing within the gel were observable at the macro, meso and nano scales. Confocal microscopy revealed that increasing shearing frequency caused inhomogeneity in pore size in our gels, indicated by higher occurrence of larger pore areas (
To improve scalability, an automated collagen mixer was developed (
Mechanical properties of tunable collagen islands
Stiffness is only one mechanical property of biomaterials that can affect cell behavior. In particular, collagen can undergo nonlinear strain stiffening in which collagen can increase in stiffness significantly under increasing applied strain. Notably, cells are able to mechanically remodel collagen to sufficiently induce strain stiffening, creating stiffness gradients over hundreds of microns (Winer et al., PloS One 2009, 4, e6382; Wang et al., Biophysical Journal, 2014, 107, 2592-2603; Rudnicki et al., Biophysical Journal, 11-20). Strain stiffening has been shown to contribute to a variety of cell behaviors including durotaxis, focal adhesion development, and higher migratory behavior. In addition, collagen is viscoelastic and can reorganize molecularly over time under applied strain leading to stress relaxation. Viscoelasticity in biomaterials has been implicated in a number of cellular processes including cell spreading, cell migration, and differentiation (Lee et al., Science Advances, 2021, 7; Lee et al., Nature Materials 2017, 16, 1243-1251; Chaudhuri et al., Nature Materials 2016, 15, 326-334; Chrisnandy et al., Nature Materials, 2022, 21, 479-487). While these properties have been studied extensively in the literature, how tissue architecture affects these mechanical properties is poorly understood.
Given the distinct architectural organization of collagen islands, the bulk mechanical properties of these gels were interrogated. Rheological shear strain sweeps revealed that all collagen island architectures maintained their strain stiffening capabilities (
Stress relaxation tests were performed at different strains to determine the viscoelastic properties of the island gels. It was found that all gels exhibit strain dependent stress relaxation properties, in which higher strains lead to faster stress relaxation (
Gel compaction assays were performed in which D1 murine mesenchymal stem cells (MSCs) were embedded in gels and monitor gel area over time (
When cell-matrix interactions in the normal or mid shear island gels were further examined, differences in dynamic protrusion activity were noticed. It was found that MSCs were highly dynamic in normal gels and demonstrated the ability to form many long protrusions and densify collagen around themselves (
Standard gel compaction assays were performed and monitored compaction over the course of 7 days (
In addition to elastic deformation of fibers within the gel, cells dynamically remodel the matrix and plastically deform the gel (Malandrino et al., PLOS Computational Biology, 2019, 15, e1006684). Plastic deformation of fibers within gels can play a major role in gel mechanics which will in turn give rise to altered cellular response. To determine the amount of plastic deformation, all cells were removed or lysed with SDS after 7 days of culture. A 1-3% increase was observed in gel area, indicating that the majority of gel compaction was non-elastic (irreversible).
Given that MSCs cultured in the mid and low shear island gels compacted less quickly than those cultured in normal gels and that MSCs cultured in island gels showed fewer dynamic protrusions, gel compaction was sought to be rescued by increasing dynamic protrusion activity. Thus, MSCs cultured in mid or low shear gels were incubated with agonists for either RhoA or all RhoGTPases. These molecular targets are known to affect cytoskeletal dynamics and are key players in collagen remodeling (Nguyen et al., Communications Biology 2022, 5, 202; Carey et al., Integrative Biology 2016, 8, 821-835). It was found that treatment with RhoGTPase agonist was sufficient to increase gel compaction rate in mid and low shear gels to that of normal gels (
To determine how island architecture modulates stem cell behavior, DI MSC's were seeded in gels that were adherent to polydopamine covered glass (
As a control to verify these results, MSCs were embedded on collagen gels of increasing density and, therefore, increasing stiffness. Indeed, we saw increased osteogenic commitment of cells in increasing collagen density (
To further investigate this differentiation behavior, localization analysis was performed for RUNX2, a key transcription factor in osteogenesis (Zhao et al., Molecular Therapy: The Journal of The American Society Of Gene Therapy, 2005, 12, 247-253; Yang et al., Nature Materials 2014, 13, 645-652). A bell-shaped curve in RUNX2 nuclear localization was again observed with the highest amount of nuclear RUNX2 found in the mid shear gels (
These results, which reveal a bell-shaped curve in osteogenic commitment in island architectures, could be explained by the complex architectures of these island gels. Though the bulk stiffness of these gels is soft, the islands provide high local stiffness which modulates stem cell behavior. This data suggests an optimal spacing of stiff inclusions to promote cell spreading. These results indicate an optimal island architecture for osteogenic differentiation which cannot be achieved in normal isotropic collagen gels.
Given that these results showed that the mid shear island gels provided optimal cell behavior in terms of spreading and differentiation, it was next investigated how this architecture could affect iPSC behavior. Single cell iPSC suspensions were encapsulated in either normal or island gels and the amount of matrigel concentration was varied. iPSCs were cultured in conditions in which matrigel made up either 20% or 50% of the final volume, and PBS was added to the 20% matrigel solutions to create equal volume solutions. In both cases, normal gels or island gels made up 50% of the final volume. Growth of iPSCs were monitored over the course of 7 days without the addition of differentiation factors (
The effect island architecture could affect stem cell fate was next explored. Localization analysis was next performed on the canonical markers for pluripotency, ZO-1 and Nanog (Simunovic et al., Nature Cell Biology, 2019, 21, 900-910). Immunofluorescence staining of these structures revealed that cells cultured in 50% island gels lose both their stemness as well as their apical basal polarity, indicated by a loss of Nanog and ZO-1, respectively (
Collagen island gels were developed which accurately depict features found in many real tissues. Island gels are relatively simple to manufacture as they only require mechanical shear to form and have tunable island size depending on mechanical shear frequency. Furthermore, these islands can be isolated from solution and resuspended in other natural ECM-derived matrices. Altogether, these island gels mimic tissue architecture, have tunable topographical and mechanical properties, and are compatible with modular assembly for tissue engineering.
The fluid dynamics model was able to obtain metrics of shear stress and energy introduced into the system. While this model does not consider key metrics such as gelation, it could give some predictive power in terms of defining the full phase space of collagen islands. The model allows for predictions on how changes in pipetting frequency, pipette geometry, and even distance of pipette from bottom of the tube could affect total energy output and, therefore, island architecture. Further studies will address how changes in these parameters will affect overall spacing and size of islands. Characterizing the phase space of these island gels will help guide optimal recipes for the assembly of mesoscopic ECM architectures and uncover key mechanical driving factors. This will allow for the study of physiologically mimicking heterogeneous and mesoscopic cues, which typical hydrogel gels do not offer but are important physiologically and pathologically (Provenzano et al., BMC Medicine, 2006, 4, 38; Cicchi et al., Journal of Biophotonics, 2010, 3, 34-43).
Bulk mechanical properties of these island gels are tunable. Their bulk stiffness can be modulated. However, AFM is still required to probe the local mechanical properties of these island architectures. In addition, while viscoelasticity has been shown to be a determinant of cell behavior in 3D hydrogels, bulk normalized viscoelasticity does not seem to be affected by introduction of island architecture. However, this constant viscoelastic behavior remains poorly understood. It is hypothesized that, while the network of the collagen has experienced significant rearrangement, longer fibers (which are the main contributors of viscoelasticity) remain unaffected. A particularly useful tool for teasing apart these mechanics are discrete fiber network models, which have been used extensively to understand local network behavior of various biomaterials under applied stresses (Mak et al., Nature Communications 2016, 7, 10323; Computational and Structural Biotechnology Journal, 2020, 18, 3969-3976). Future studies will incorporate these models to better understand the mechanical contributions of these islands to bulk properties. Overall, these data suggests that local stiffness, island spacing, and connections between islands are all important cues that affect cell behaviors in these gels.
MDA-MB-231s (Lifeact GFP) were a gift from the Lauffenburger lab. They were cultured in DMEM (Gibco™11965092) with 10% FBS (Gibco™16000-044) supplemented with 1% penicillin-streptomycin (5,000U/ml, Gibco™15070-063). Normal human lung fibroblasts (ATCC, PCS-201-013) were cultured in Lonza FGM-2 BulletKit (CC-3132) or RPMI1640(Gibco™11875-093) with 10% FBS and 1% Pen-Strep. Cells were all maintained at 37.0 and 5% CO2.
D1 MSC cells were obtained from the American Type Culture Collection (ATCC; Manassas, VA, USA). ATCC validated all cell lines by Short Tandem Repeat Analysis. Cells were maintained at 37° C., 5% CO2. D1 MSCs were maintained in Dulbecco's Modified Eagle's Medium (DMEM) medium supplemented with 10% fetal bovine serum, 1% L-glutamine, and 1% penicillin/streptomycin. Media was changed every other day, and cells were split every 3-4 days. To generate MSC cells expressing LifeAct-GFP, lentiviral particles containing a pLentiCMV-MCS-LifeAct-GFP vector were used. Cells were selected with puromycin (10 μg ml−1) and were cultured at no more than 80% confluency at no greater than passage 25 in serum-supplemented DMEM.
The hiPSCs were a gift from Dr. Stuart Campbell (Department of Biomedical Engineering, Yale University). hiPSCs were cultured on TC-treated 6-well plates (Costar, Corning) coated with lactose dehydrogenase elevating virus (LDEV)-free human-embryonic-stem-cell-qualified Matrigel (Corning, 354277) in mTeSR1 media (STEMCELL Technologies) at 37° C. in 5% CO2. Media was changed daily. hiPSCs cultured in mTeSR1 were passaged at 70% confluency as pluripotent aggregates with manual selection using ReLeSR (STEMCELL Technologies) following the manufacturer's protocol.
Each collagen gel was made by adding sufficient 0.5 N NaOH to neutralize a mixture of double-distilled H2O, 10×PBS, and acetic-acid-solubilized type I rat tail collagen (Corning, Corning, NY, USA) on ice for a final collagen concentration of 2 mg/ml. Island gels were made by first neutralizing 350u1 of collagen. Stained collagen gels were made by incorporating 10% 647-ester dye (ThermoFisher) stained collagen. The gels were then allowed to sit at room temperature for 6.5 minutes. Finally, the gels were sheared with a p200 pipette tip set to 175u1 at a specified frequency for 3 minutes before being plated. Prior to depositing the collagen, plates used for collagen gel seeding were surface-coated with polydopamine, as previously described (Park et al., ACS Applied Material Interfaces 2019, 11, 23919-23925; Lee et al., Science 2007, 318, 426-430; Nguyen et al., Communications Biology, 2002, 5), which allowed collagen gels to stick to the surface of the plates to prevent detachment of the gel. After plating, gels were transferred to an incubator at 37° C. with 5% CO2 for 1 hour.
hiPSCs at 70% confluency or less were passaged as single cells using Accutase (STEMCELL Technologies), washed in mTeSR1 media, and resuspended as a single-cell suspension in mTeSR1 media with 10×10-6 m ROCK inhibitor (Y-27632, STEMCELL Technologies) to prevent dissociation-induced apoptosis. For 50% collagen gels, isotropic or island gels were made and mixed in equal parts on ice with Matrigel (Corning, 354277). For 20% matrigel:collagen gels, isotropic or island gels were made and mixed with Matrigel and PBS on ice such that the resulting gel was 20% Matrigel by volume. Collagen gels were made, and an appropriate number of cells were mixed to achieve a final concentration of 100000 cells/ml. After gelation, mTeSR1 media with 10×10−6 m ROCK inhibitor was added. 24 h post encapsulation, the media was changed to mTeSR1 which was replenished daily.
Encapsulation of MSCs within hydrogels
D1 MSCs or D1 MSC-LifeActRFP cells were mixed with the collagen solution prior to gelation to achieve a final cell concentration of 15000 cells/ml. Cells were cultured in 50%/50% mixture of the osteogenic induction medium and adipogenic induction medium (differentiation medium). The differentiation medium contained 50 μg/ml L-ascorbic acid (MilliporeSigma), 10 mM β-glycerophosphate (MilliporeSigma), and 0.1 μM dexamethasone (MilliporeSigma) in the DMEM growth medium.
MSC time lapses were performed in 12 well glass bottom dishes which had been polydopamine coated. 12 hour time lapses were taken on Day 1 and Day 6. Automatic cell segmentation was obtained using Imaris (Oxford Instruments). Manual filtering was done to determine which cells were to be used for downstream analysis. From these segmentations, cell movement was tracked and metrics such as mean speed, sphericity, and migration track length over the timelapse were obtained. Mean speed is defined as the average of instantaneous speeds between timepoints.
Custom Python3 scripts were used to calculate MSD plots. Trajectories were exported to Python to calculate average speed and mean squared displacement. Average speed specifically refers to the mean of the absolute value of the net displacement of the cell center per hour. Mean squared displacements MSD(n) were computed with the following equation:
where N is the total number of steps, n is the n-th step, and x,y,z are the x,y,z coordinates, respectively.
Gel Compaction
Gel compaction experiments were performed in 24 well plastic bottom plates. Wells were incubated with 3% BSA for 1 hour and then washed with PBS to avoid adhesion of the gel to the plate. D1 MSCs were mixed with 500 μL of collagen solution prior to gelation to achieve a final cell concentration of 600000 cells/ml. Cells were cultured in the differentiation medium described above, and media was changed every day. Images of the gels were taken every day for 7 days. After 7 days of culture, gels were treated with an 8% Triton and 0.125% Trypsin-EDTA solution for 30 minutes to decellularize gels and observe the degree of plastic deformation. Images of the gels were taken after decellularization. Timelapses of gel compaction assays were performed in 15 well plates (Ibidi). Cells were incubated with 3% BSA for 1 hour and then washed with PBS to avoid adhesion of the gel to the plate. D1 MSCs were mixed with 15 μL of collagen solution prior to gelation to achieve a final cell concentration of 600000 cells/ml. Cells were cultured in 50 μL of the differentiation medium described above. Images were taken every 6 minutes for 12 hours.
MCF10As (ATCC, CRL-10317™) were cultured with MCF10A growth medium which includes DMEM/F12 (Gibco™, 11330-032) supplemented with 5% horse serum (Gibco™16050122), 20 ng/ml EGF, 0.5 mg/ml hydrocortisone (Sigma H- 0888), 100 ng/ml cholera toxin (Sigma C-8052), 10 ug/ml insulin (Sigma 1-1882) and 1% penicillin-streptomycin (5,000U/ml, Gibco™15070-063). To produce MCF10A acini, 24 well plates were first coated with a thin layer of pre-thawed Matrigel (Corning 354230) and then kept in incubator for 30 minutes to allow Matrigel gelation. Around 5K to 10K MCF10A single cells were then seeded on top of the Matrigel and maintained with MCF10A assay medium in 37.C, 5% CO2 incubator until day4. MCF10A assay medium has the same components with MCF10A growth medium but with 2% horse serum and no EGF. MCF10A maintenance and acini formation protocols are adopted from the Brugge lab. Cell recovery solution (Corning354253) was used to extract MCF10A acini from Matrigel following manufacturer's protocol. Briefly, MCF10A acini (in Matrigel) were first washed with ice cold PBS 3 times followed by 1 hr ice incubation in cell recovery solution. Then MCF10A acini were spun down and washed in ice cold PBS for 3 times at 70 g for 2 minutes in pre-chilled centrifuge. Washed MCF10A acini were seeded in normal or thick collagen for further study.
Acid solubilized collagen I (Corning354249) was first neutralized with NaOH and then diluted with suspended cells (or acini) to a final concentration of 2 mg/ml or 1 mg/ml (MCF10A study) on ice. Culture media were added after 1 hr gelation at 37° C. For labeled ECM studies, the initial collagen solution was labeled with Alexa Fluor 647 NHS Ester (Succinimidyl Ester) and dialyzed as before (Debnath et al., Methods, 2003, 30, 256-268). To note, the surface of the multi-well plates used for imaging were all pre-coated with polydopamine to anchor the collagen gel (Lee et al., Science, 2007, 318, 426-430; Yu et al., RSC Advances, 2014, 4, 7185), including thick collagen gels.
Acid solubilized collagen I (Corning354249) was neutralized with NaOH in the same way as in normal collagen preparation mentioned above (step 1 in
To further validate the replicability of the unique Collagen I architecture, a computer-controlled extruder was designed and constructed. Conventional off the shelf syringe pumps offer the accuracy required for this application, however, they tend to be heavy, bulky, costly and few offer the option of running custom pumping sequences without bypassing the whole controller board and connecting directly to the motor. It was therefore decided that to achieve the desired pumping in/out cycle, the best option would be to design a linear actuator that could operate any syringe and would be light enough to attach to a scaffold over the hot plate. The main components of the actuator are a 3D-printed frame, a Nema 17 stepper motor, a A4988 stepper driver, a set of 20T/60T GT2 gears, a T8 2 mm pitch lead screw, and an Arduino Uno microcontroller. The complete structure was printed in about 11 h using an Elegoo Saturn resin printer. The Arduino IDE was used to program and upload the code on the board. Using the known gear ratios and rotational-to-linear motion conversion of the lead screw, the speed of the stepper motor was adjusted to obtain the desired flow rates. The complete collagen shearing system can be built for less than $200.
Drugs were reconstituted and stored following manufacturers' recommendations and diluted in culture media to working concentrations. Specifically, 20 μM pan-MMPs inhibitor GM6001(Abcam, ab120845), 30 μM ROCK inhibitor Y27632 (Abcam, ab144494), 10 μM Cdc42 inhibitor ML141 (Calbiochem®217708), 50 μM formin inhibitor SMIFH2 (Abcam, ab218296), 50 μM ARP2/3 inhibitor CK666 (Abcam, ab141231), 100 μM Rac1 inhibitor NSC23766, 20 μM Rac1 inhibitor EHT1864 (Cayman 17258) were used. Fresh media mixed with specific drug were prepared right before experiment and replaced daily. In the drug assays, at least two independent experiments were performed with at least two replicates (technical duplicates) in each experiment.
The microplate reader was first pre-heated to 37° C. For normal collagen measurement, acid solubilized collagen was neutralized and diluted to 2 mg/ml as above-mentioned. Ice-cold collagen solution was aliquoted into 96 well plates (80 μl per well) and absorbance at 405 nm (A405) was tracked every minute for 20 minutes. For thick collagen measurement, after following the standard thick collagen protocol to step 4 (without ice incubation), thick collagen solution was aliquoted into 96 well plates (80 μl per well) and readouts of A405 were recorded every minute for 20 minutes.
Culture media were first removed and then the gels were rinsed 3X gently with 1×PBS. Cells were then fixed with 4% formaldehyde for 30 min and permeabilized by 0.3% Triton X-100 for another 30 min. After fixation and permeabilization, cells were blocked with 1% BSA followed by primary antibody incubation (1:200), 3×PBS wash and secondary antibody incubation (1:500) supplemented by Hoechst stain (1:2000) and Alexa Fluor™647 Phalloidin (Invitrogen™A22287, 1:200). Fluorescence imaging were acquired within one week after immunostaining.
Gels were rinsed twice with PBS and fixed with 4% paraformaldehyde (Thermo Fisher Scientific) in PBS for 25 min at room temperature. Following fixation, gels were permeabilized with 0.2% Triton X-100 in PBS for 1 h at room temperature. They were then blocked with 3% BSA in PBS containing 0.01% Triton X-100 for 3 h at room temperature. The samples were then incubated for 24 hat 4° C. with the primary antibody Nanog (1:100; α-Rabbit; Cell Signaling Technologies), SOX2 (1:100; α-Rabbit; Cell Signaling Technologies), Oct4 (1:200; a-Rabbit; Cell Signaling Technologies), Vimentin (1:200; α-mouse; Santa Cruz), SNAIL (1:200; α-Goat; R&D Systems), Brachyury (1:100; a-Goat; R&D Systems), or RUNX2 (1:100; Mouse; Cell Signaling Technologies). After washing for 3-5 h at room temperature, samples were incubated overnight at 4° C. with secondary antibody Alexa 647 goat-a-rabbit (1:1000 in PBS; Invitrogen), Alexa Fluor donkey-α-goat (1:1000 in PBS; Invitrogen), or Alexa 488 goat-α-mouse (1:1000 in PBS; Invitrogen), DAPI (1:2,000, Invitrogen) and phalloidin-Alexa 555 (1:500; Abcam). After at least 3 hours of washing, samples were mounted in Vectashield (Vector Laboratories) before imaging.
ALP was stained with a FastBlue working solution of 500 μg/ml Fast Blue BB (MilliporeSigma) and 500 μg/ml naphthol-AS-MX (MilliporeSigma) phosphate in an alkaline buffer (100 mM Tris-HCl, 100 mM NaCl, 0.1% Tween-20, 50 mM MgCl2, pH=8.2). Fixed samples were first washed three times in DPBS, equilibrated in alkaline buffer for 15 min, then incubated in FastBlue working solution for 60 min at room temperature. The samples were then washed in alkaline buffer for 15 min followed by 15 min in DPBS. For Oil Red staining, gels were equilibrated with 60% isopropanol for 30 minutes before being incubated in Oil Red in 60% isopropanol for 1 hour. Gels were then washed three times with water for 1 hour before imaging. For Alizarin Red staining, fixed samples were first equilibrated with water for 30 minutes. Gels were then incubated in 2% ARS solution (Sigma) for 30 minutes. Gels were then washed three times with H2O for an hour. Samples were then imaged after 7 days on a Leica DMH transmitted light microscope.
To quantify stem cell differentiation, mRNA from iPSCs cultured in gels for 7 days were extracted. At least 3 gels per replicate were suspended in Trizol and homogenized with a 20G needle. RNA was isolated by phenol-chloroform extraction and subsequent RNA extraction columns from the RNEasy Extraction Kit (Qiagen). 5 μg of RNA were reverse transcribed and amplified using the iTaq Universal SYBR Green One Step Kit (BioRad). Samples were analyzed using the Bio-Rad iTaq Universal Probes One-Step Kit in 20-μl reactions run at 50° C. for 10 min and 95° C. for 1 min, followed by 40 cycles of 95° C. for 10 s and 60° C. for 2 minutes per the manufacturer's recommendations. Reactions were performed on an Applied Biosystems 7500 instrument.
Imaging was performed using a Leica SP8 confocal microscope. Time-lapse imaging (XYZT mode) was taken with a 20× objective (0.75 NA), with time interval set to 3 minutes or 6 minutes and z-step size set to 2 μm. Day0 time-lapse imaging was acquired between around 0hr to 12 hrs post embedding cells into gels. Day1 time-lapse imaging was acquired between around 24 hrs to 36 hrs post gel-embedment. Overview tilescan images were taken at day1, day2 or day5 respectively.
A Leica SP8 laser scanning confocal microscope with a x10 objective or x 5 objective (Wetzlar, Germany) was used for live imaging of MSC's. The 10x objective was used for 7 day time lapse experiments, and Z-stacks were taken with a thickness of 2 um. 12 hour time lapses were taken on Day 1 and Day 6. The 10x was used for 12 hour gel compaction experiments, and Z-stacks were taken with a thickness of Sum. A temperature of 37° C. and a 5% CO2 atmosphere were maintained using a humidified OKO labs live-cell imaging incubator. 3D fluorescence images were taken with either a 20x objective or 40x water objective. 3D Z stacks were taken with a thickness of 2 um. Cell and nuclear outlines were manually traced in ImageJ.
To track cell migration, XYZT data were first reduced in dimensions by conducting standard deviation projection along the z-axis. The XYT data were then processed by TrackMate in Fiji and produced cell trajectories. Trajectories were exported to MATLAB to calculate average speed and mean squared displacement. Average speed specifically refers to the mean of the absolute value of the net displacement of the cell center per hour. Mean squared displacements were computed with the following equation (Gorelik, Nature Protocols 2014, 9, 1931-1943):
In this equation, N is the total step number, n is the n-th step, x,y are the x,y coordinates respectively. For morphology analysis, z projected tilescan images were segmented and analyzed by in-house MATLAB codes. Segmentation performance was validated, with wrong segmentation manually corrected or removed. For difficult automatic segmentation data such as the Y27632 treated condition, manually tracked data were supplemented. The definitions of elongation, compactness, sphericity and shape variance were described in
An Anton Parr Shear Stress Rheometer was used for mechanical characterization of collagen gels with the 25-mm parallel-plate geometry and a 500 μm gap. To prevent slip between the gel and the plate, no. 1 25-mm cover glasses (VWR) used for the top plate and a 40 mm cover glass were chemically treated with polydopamine and attached to each plate of the rheometer with double-sided tape (3M 666). The rheometer was then zeroed and calibrated. The temperature of the system is preset and maintained at 37° C. Around 300 μL collagen was pipetted onto the rheometer, the top plate was quickly lowered 500 μm, and the sample was kept in a custom-made humidity chamber to prevent evaporation. After at least 60 minutes of gelation, gels were immersed in PBS and allowed to sit for at least 15 minutes before taking measurements. Then, the shear modulus was measured at 2% strain and at 0.1 Hz. The shear modulus was determined and analyzed using custom Python scripts. Frequency sweeps were carried out at 2% strain.
In the testing of hydrogels, collagen was deposited onto the bottom plate of the rheometer immediately before gelation, and the top plate was lowered rapidly so that the gel formed a uniform disk between the two plates. Approximately, 350 μL collagen was pipetted onto the rheometer, the gap was set to 500 μm, and the sample was kept in a custom made humidity chamber to prevent evaporation. Polymerization progress was monitored by imposing three cycles of 0.5% strain every 5 min, measuring the shear storage modulus G′ as a function of polymerization time. After at least 60 minutes of gelation, gels were immersed in PBS and allowed to sit for at least 15 minutes before taking measurements. For strain sweep measurements, collagen gels were subjected to 5 oscillations at 0.1 Hz at increasing amplitudes from 2 to 12% in 2% increments and 12 to 100% in 4% increments. Custom Python scripts were used to determine the differential shear moduli and subsequent metrics. For stress relaxation measurements, strains were applied with a rise time of 0.15 s. Only one stress relaxation test is conducted on any given sample. Custom Python scripts were used to determine the relaxation moduli, peak stress, and half max relaxation time (t).
The collagen sample is placed in 4% PFA for 1 hour at room temperature. The sample is then soaked twice in PBS for 10 minutes each. This is followed by two 10-minute ddH2O washes. The sample is then put through a graded ethanol+H2O wash for 10 minutes each in this order: 30% EtOH, 50% EtOH, 66% EtOH, and 100% EtOH. The sample was then put through a graded ethanol +HMDS wash for 10 minutes each in this order: 30% HMDS, 50% HMDS, 66% HMDS, and 100% HMDS. Samples were then placed on aluminum foil and allowed to dry in the fume hood overnight. Subsequently, the samples were mounted on a support with carbon tape and covered with an 8 nm layer of iridium with a sputter coater. The samples were then imaged with a scanning electron microscope.
For hydrogels, the collagen sample is placed in 4% paraformaldehyde (Thermo Fisher Scientific) for 1 hour at room temperature. The sample is then washed twice in PBS for 10 minutes each. This is followed by two 10-minute ddH2O washes. The sample was then through a graded ethanol+H2O wash for 10 minutes each in this order: 30% EtOH, 50% EtOH, 66%
EtOH, and 100% EtOH. The sample was then put through a graded ethanol+HMDS wash for 10 minutes each in this order: 30% HMDS, 50% HMDS, 66% HMDS, and 100% HMDS. Samples were then placed on aluminum foil and allowed to dry in the fume hood overnight. Subsequently, the samples were mounted on a support with carbon tape and covered with a 8 nm layer of iridium with a sputter coater. The samples were then imaged with a scanning electron microscope.
Sampling and statistical analyses of various results plots are indicated in their corresponding figure caption. For shear modulus, unpaired t-test is performed. For statistical comparisons in drug studies, one-way ANOVA with post-hoc Tukey HSD test was performed. * indicates p-values<0.05. ** indicates p-value<0.01. *** indicates p-value<0.001, **** indicates p-value<0.0001. # indicates current condition is significantly different (p<0.05) with every other condition unless otherwise annotated.
GraphPad Prism was used for all statistical analyses. In vitro experiments were repeated at least three independent times. To compare differences between more than two groups, a one-way ANOVA with Tukey's post-hoc test was used. Different levels of statistical significance were set at *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001.
HCC cases were selected from the Yale New Haven Hospital pathology database. No approval from a research ethics committee was required for this study, as coded tissue obtained from routine diagnostic workflow was used and the included patients are not affected by the study. Anonymous or coded use of redundant tissue for research purposes is part of the standard treatment agreement with patients, to which patients may opt out. None of the included patients submitted an objection against use of residual material.
Antigen retrieval and slide staining for SHG imaging were adapted from previous protocols. Formalin fixed slides were first incubated at 55° C. for 10 minutes. Slides were then immersed in Xylene baths 2 times, 5 min each time, to wash off paraffin. Slides were then immersed in a series of ethanol baths to rehydrate tissue. Slides were then soaked in DI water to hydrate. Tissues were then immersed in 10 mM Sodium Citrate, 0.05% Tween 20, pH 6 solution at 95° C. for 15 minutes. The solution was then brought down to room temperature, and slides were removed and carefully dried. The sample was then outlined with a hydrophobic pen. The sample was then soaked in TB S+0.025% TritonX-100 two times for 5 minutes each. The tissue was then blocked in TBS+10% BSA at RT for 2 hours. Tissue samples were then stained with DAPI for 1 hour at room temperature. Samples were mounted in Vectashield before imaging.
Images were acquired using two-photon microscopy (MaiTai Ti:Sapphire Laser, Spectra Physics) with a ×40 and excitation at 890 and 1,090 nm.
Flow simulations corresponding to the pipetting cycles were performed in Comsol Multiphysics. The computational domain was originally limited to the pipette tip, however due to potentially significant interactions between the flow and the microcentrifuge tube, the domain was expanded to include the fluid both inside the micropipette tip and the microcentrifuge tube. The geometry of the P200 micropipette (Thermo Scientific ART) and the 1.5 mL microcentrifuge tube (USA Scientific) were built in Comsol Multiphysics as an axisymmetric model. The model was meshed using the adaptive mesh refinement tool within the Time-Dependent Solver to obtain a grid of 5716 triangular elements and 438 edge elements. The effects of Col. I crosslinking and other chemical reactions were ignored, so the properties of the fluid domain were simplified as water at 303.15 K. To simulate the cycle of pipetting in and pipetting out at different frequencies, the inlet boundary condition of the Laminar Flow study was defined by overlapping two functions: a positive mass flow rate (m′) times a step function going from 0 to T seconds, and a negative mass flow rate (−m′) times a step function from T to 2T seconds. Periods (T) of 2, 5 and 10 seconds were defined in a Parametric Sweep study. To reduce computational time, a single cycle of flow out-in was simulated for each condition. Thus, the time length of each Time Dependent study was set at 2T (0.1 sec steps) and solved using the PARDISO Direct Solver. Using the solution for shear rate ({dot over (γ)}), the shear stress (r) was computed for each element through the geometry using for each element through the geometry where μ is the viscosity of the fluid. The computed shear stress was then integrated over the volume of the geometry V to obtain the instantaneous shearing energy for each time step n:
ε[n]=∫vτ[n]dV
which is the average energy between two consecutive timepoints. Therefore, for any length of time T, the total shearing energy delivered to the fluid can be computed as
where N is the total number of time steps between times 0 and T.
The disclosures of each and every patent, patent application, and publication cited herein are hereby incorporated herein by reference in their entirety. While this invention has been disclosed with reference to specific embodiments, it is apparent that other embodiments and variations of this invention may be devised by others skilled in the art without departing from the true spirit and scope of the invention. The appended claims are intended to be construed to include all such embodiments and equivalent variations.
This application claims priority to and the benefit of U.S. Provisional Application No. 63/481,511, filed Jan. 25, 2023, and U.S. Provisional Application No. 63/382,571, filed Nov. 7, 2022, the disclosures of which are incorporated herein by reference herewith in their entireties.
This invention was made with government support under R35GM142875 awarded by the National Institutes of Health (NIH). The government has certain rights in the invention.
| Number | Date | Country | |
|---|---|---|---|
| 63481511 | Jan 2023 | US | |
| 63382571 | Nov 2022 | US |
