EP4164706A1 - Device and process for tissue-engineering and regenerative medicine - Google Patents

Abstract

a process and device for direct in-vitro to in-vivo transfer of a living microcarrier culture by controlled dehydration into a viscoelastic material capable of cell-protection during delivery. One particular purpose of the present invention is that of providing a process and device for direct in-vitro to in-vivo transfer of living microcarrier co-cultures.

Description

Device and process for tissue-engineering and regenerative medicine

Technical field

This invention generally pertains to the tissue engineering and regenerative medicine fields.

Introduction and Background

Biomaterials are increasingly used for in-vivo cell delivery (some examples: US2017136153A1, W02012149358 Al, W02013036875A1), where they often take the name of scaffolds(Garg et al., 2012). Besides offering adhesion sites for anchorage- dependent cells, they allow additional functionalities such as defined co-culture or pre vascularization (W02008008229A2). An interesting alternative is also to deliver cells and scaffolds separately. This is possible for mobile cells such as hematopoietic stem cells, when strong homing factors are known (W02017/136837A1). Scaffolds can further be composed of a variety of synthetic or natural polymers, but also decellularized extracellular matrix (for example, W02017/223529, W02016168752 Al).

Injectable scaffolds, for example preformed large scaffolds(Bencherif et al., 2012a), W02012149358 Al, but also other injectables, such as viscoelastic hydrogels (Nih et al., 2017) or cements (for example W02014160232A2) are of particular utility since they permit minimally invasive delivery. For scaffold injection, it is sometime necessary to dehydrate scaffolds; this is typically obtained by applying some sort of mechanical pressure during injection (Bencherif et al., 2012a) and/or partial dehydration before actual injection (Beduer et al., 2015a).

Microcarriers are particles on the scale of tens of micrometers to the millimeter range. They are intended for the culture of adherent cells in suspension. Many different microcarriers are commercially available (the Cytodex, Cytopore and Cultisphere series being among the more renowned), and many fabrication methods and compositions established in the scientific and patent literature (example: EP1801122A1; defined shapes in W02014037862A1). Microcarrier-based cell culture is also a well-established art. The typical application of microcarriers is large-scale production of cells or other biological such as antibodies or cells (example: hematopoietic stem cells, W00046349A1 and W02012127320A1); this can elegantly be combined with cell harvest by affinity to microcarriers (W02009/002456 A2). Microcarriers are also increasingly used to deliver cell therapy(example: US2015361395A1), since they can provide cells with adhesion during the transplantation process (Newland et al., 2015).

Despite the enormous potential that microcarrier culture has in terms of cell amplification, screening applications, ease of use and robustness, there is a relative lack of methods to go directly from a microcarrier culture to a dense in-vivo injectable. This invention aims to address this shortcoming of the current state of the art. The necessity for a defined transition with substantial volume reduction from a dilute suspension to a dense, transplantable injectable is particularly a problem for cells expanded to high numbers (typically, various stem cells such as embryonic stem cells, induced pluripotent stem cells, or endogenous stem cells such as hematopoietic stem cells and lymphoid progeny or neural stem cells), but also for postmitotic cells with a high-demand metabolism (for example, transient amplifying or progenitor populations, neurons, transport epithelia, cardiomyocytes or immune cells).

The challenge in transplantation of a microcarrier culture is primarily its dilute nature. At present, washing and recovery of microcarriers is routinely performed, typically by sedimentation (Croughan et al., 2016). However, at best, this allows to recover the microcarriers with their equilibrium amount of fluid associated, a possible option to obtain some improvements in mechanical properties being the addition of thickeners (for example, US 5,980,888; W001/12247, EP3210634A1). To avoid additional reagents, denser suspensions with partial compression of the microcarriers are typically produced from dry material (Xia et al., 2017). For this, one can for example use lyophilized powder as supplied commercially for many microcarriers, or lyophilize custom-made microcarriers. The advantage is of such suspensions under compression is that substantial forces can be sustained, such that finite mechanical properties are obtained(Xia et al., 2017). It is however difficult at present to obtain such partially compressed suspensions from dilute cultures.

The reasons for the difficulties are rooted in the need to remove pore fluid while preserving cell presence, viability and in some models such as co-cultures, also the cell cell relations. It is an object of this invention to address the problem of dehydration of dilute microcarrier cultures to dense, partially compressed paste-like materials with tissue-like mechanical properties, while preserving cell viability, presence and interactions, if applicable, in a reproducible way. This in turn enables applications where in a first step, microcarriers are cultured in large, dilute volumes to satisfy high metabolic needs arising from stem and progenitor cells, neurons, cardiomyocytes, transport epithelial cells or other highly active cells, and where in a second step injectable implants with high mechanical stability arising from partial microcarrier compression need to be produced. The application is to simultaneously transplant the cells and form mechanically stable local niche.

During large-scale microcarrier dehydration, two parameters should be controlled independently:

First, the aspiration pressure applied to the interstitial fluid. Together with an empirical calibration curve characteristic for each material, this determines the final microcarrier density and indirectly the mechanical properties of the final implant. Second, the dehydration rate, as expressed by the fluid flow rate through relevant portions or all of the microcarrier material being compacted. As cells are sensitive to shear(Tanzeglock et al., 2009), this must be controlled if cell presence, integrity and if applicable cell interrelations are to be preserved. Various filter systems exist in the art, including sterile filter systems for automated stem cell harvest under gentle and controlled conditions (US2005/0048036 Al, W02006014158A1), but the difficulty of concentrating a microcarrier suspension for delivery into a patient, animal or even in 3D printing applications to a cell culture system are particular. First, the pressure and especially the flow rate need to be controlled to values that are very low compared with the usual target of obtaining rapid fluid removal in filtration systems, otherwise the cells will be flushed from the microcarriers (this phenonenon has been exploited for recovery of hematopoietic stem cells from microcarriers, see W02013141731A2 p. 16, 1. 21ff.). Second, the compacted microcarriers need to remain with either directly a suitable syringe or a transfer device, rather than sticking to the fluid removal device, since otherwise it needs to be scraped of a membrane with concomitant risk of loss of cell viability, and generation of foreign particles. The present invention addresses these challenges.

While many microcarriers lend themselves to the present invention, there are differences in their properties that make their ease of use or performance different. Scaffolds with hard, spherical particles yield relatively brittle pastes that are nearly unable to sustain shapes other than externally imposed ones; such an example is provided in the literature by partial rehydration for culture of metabolically relatively inactive cells (Fig. 8h in (Xia et al., 2017)), but can also be reached from suspension culture with the present invention by using for example cytodex microcarriers. In this case, the primary advantage of dehydration lies in the cell density. Softer, porous materials, for example the commercial material cultisphere G or possibly newer non-porous (EP1801122A1) or porous gelatin carriers (EP3210634A1) make for more stable biomaterial pastes, most likely due to better shape-adaption and particle interlocking. The invention is particularly useful in conjunction with microcarriers exhibiting a plateau in compression such as the ones describes in W02017/029633; these are able to maintain their mechanical properties even for major volume changes; once adjusted these are resilient to loss of pore fluid by evaporation or dilution by addition of small to moderate amounts of liquid, for instance by contact with body fluids.

Scaffolds have also been used to provide stromal functionality, for example by allowing implantation of mesenchymal stem cells for nerve repair (CN104491925B), or for co culture and co-transplantation of stem cells with stromal components, for instance to mimick a bone marrow environment for acute myeloid leukemia cancer stem cells, hematopoietic stem cells or other stem cells along with a mesenchymal stem cells cultured on the scaffold to provide stromal support (US2011/0207166A1). The present invention can also be advantageously used for such applications, provided the co-culture can be implemented on microcarriers suitable for this invention. An example regarding such an application is provided here, where particles as described in W02017/029633 are coated with collagen 1 for compatibilization with the mesenchymal OP9 cell line, establishing an expansion co-culture of hematopoietic stem cells on the collagen 1-OP9 coated microcarriers. After compaction by the methods of the present invention, the co culture forms an injectable, ready for subcutaneous or intraosseous implantation by minimally invasive delivery (injection).

A possible application of the present invention is the generation of an ectopic or new marrow niche for hematopoietic stem cells by means of a co-transplantation with stromal cells on microcarriers. It is indeed the basis of hematopoietic stem cell transplantation that hematopoietic stem cells are exquisitely dependent on their niche for proliferation and maintenance. In fact, to guarantee successful engraftment, at least partial emptying of the hematopoietic niche of the recipient is at present necessary. Chemotherapy and whole-body irradiation are the major means to this end, although niche manipulation by injection of supplementary mesenchymal stem cells (W02016151476A1) together with the use of mobilizing agents such as GM-CSF are also being explored. An alternative means is to provide a local, possibly ectopic niche altogether (W02017136837A1, filled by separate transplantation of hematopoietic stem cells). The teachings of this invention can be used to create an injectable, yet coherent and shape-stable implant from a microcarrier-based co-culture of stromal cells (such as the OP9 line) and hematopoietic stem/progenitor cells after efficient in-vitro expansion under microcarrier culture conditions (cytokine-based, see for example W02017075389A1, or based solely on the action of the OP-9 or other stromal cells).

Brief summary of the invention

In order to address and overcome at least some of the above-mentioned drawbacks of the prior art solutions, the present inventors developed a process and device for direct in- vitro to in-vivo transfer of a living microcarrier culture by controlled dehydration into a viscoelastic material capable of cell-protection during delivery. One particular purpose of the present invention is that of providing a process and device for direct in-vitro to in- vivo transfer of living microcarrier co-cultures.

The invention as described in here serves to compact a microcarrier culture or co-culture into an injectable implant with a paste-like consistency. The final implant remains injectable, but due to partial dehydration, has some mechanical strength, as evident for instance by rheometric G’ values in the range of lOPa to IMPa, more preferentially lOOPa to lOkP and most preferentially 500Pa to 5kPa.

The partial dehydration system consists of a fluid drain capable of absorbing amounts at least equal, but typically by far (lOx or more) exceeding the amount of fluid to be absorbed from the microcarrier culture. It further is capable of separately, and stringently, regulate both final interstitial fluid pressure to be achieved (between 20 Pa and lOkPa, preferentially 50 Pa to lkPa, and even more preferentially between 100 Pa and 800 Pa below atmospheric pressure), and the maximum fluid flow rate experienced by microcarriers to subsequently be delivered in-vivo (or for in-vitro applications such as 3D printing) in the range between O.Olmm/s and lOcm/s, more preferentially between O.lmm/s and 5cm/s, and even more preferentially between 0.5mm/s and lOmm/s. In a preferred embodiment, the invention further comprises a transfer recipient. In this preferred embodiment, the dehydration process is initiated when the transfer device is loaded with the microcarrier culture, optionally through a filling column. The flow rate can be imposed by the dehydration system itself, or optionally, by a self-assembling dense plug of microcarrier material at the orifice of the transfer device in contact with the dehydration device. In this case, the dimensions of the orifice are key for the dehydration process. After the dehydration to the desired final pressure (which can optionally be monitored, for example by a pressure gauge or glass capillary in touch with the material being dehydrated), the compacted, paste-like microcarrier culture (or co-culture) is confined in the transfer device and can be manipulated or transported. The transfer device typically ensures connection to an injection syringe, or is itself an adapted syringe (with an air vent for piston insertion), and also ensures connection to the delivery tubing (typically, a catheter or blunt needle), through which the compacted, living microcarrier culture or co-culture is then delivered in-vivo. It is optionally possible to sub-sample some the compacted material to ensure appropriate composition and viability prior to injection.

The above and other objects, features and advantages of the herein presented subject- matter will become more apparent from a study of the following description with reference to the attached figures showing some preferred aspects of said subject-matter. Detailed description of the invention DEFINITIONS

For the sake of clarity, some definitions will be provided hereinafter. As used herein, a “scaffold” is any three dimensional material having a framework architecture, for instance a support structure comprising hollow spaces within it. In preferred embodiments, a scaffold is an artificial structure capable of supporting cell culture and/or three- dimensional tissue/organ formation in vivo, in vitro or ex vivo. In this context, a scaffold is also referred herewith as a “biomaterial” or “bioscaffold”. In this embodiments, a scaffold can be considered the physical structure (including biodegradable and/or permanent materials) upon which or into which cells directly or indirectly associate or attach. A bioscaffold may allow or facilitate cell attachment and migration, delivers and retains cells and biochemical factors, enables diffusion of vital cell nutrients and expressed products, exerts certain mechanical and biological influences to modify the behaviour of the cell phase and so forth.

As used herein, a “polymeric material” is any material comprising polymers, large molecules (also known as macromolecules) composed of many repeated smaller units, or subunits, called monomers, tightly bonded together by covalent bonds. As used herein, the term “gel” refers to a non-fluid colloidal network or polymer network that is expanded throughout its whole volume by a fluid. A gel is a solid three-dimensional network that spans the volume of a liquid medium and ensnares it through surface tension effects. The internal network structure may result from physical bonds (physical gels) or chemical bonds (chemical gels) such as covalent, ionic, hydrogen and/or Van der Waals bonds.

As used herein, a “microcarrier” is a particle on the scale of few micrometers up to millimeters, intended for the culture of adherent cells in suspension. Microcarriers can comprise or be substantially composed of polymeric materials such as plastic materials, biopolymers and/or ceramic materials. Many different microcarriers are commercially available (the Cytodex, Cytopore and Cultisphere series being among the more renowned), and many fabrication methods and compositions established in the scientific and patent literature (example: EP1801122A1). Microcarrier-based cell culture is also a well-established art. The typical application of microcarriers is large-scale production of cells or other biological such as antibodies (example: hematopoietic stem cells, W00046349A1); this can elegantly be combined with cell harvest by affinity to microcarriers (W02009/002456 A2). Microcarriers are also increasingly used to deliver cell therapy (example: US2015361395A1), since they can provide cells with adhesion during the transplantation process (Newland et al., 2015).

In some embodiments, microcarriers according to the invention are substantially composed out of hydrogel. As used herein, the term “hydrogel” refers to a gel in which the swelling agent is water. A hydrogel is a macromolecular polymer gel constructed of a network of cross-linked polymer chains. It is synthesized from hydrophilic monomers, sometimes found as a colloidal gel in which water is the dispersion medium. Hydrogels are highly absorbent (they can contain up to over 90% water) natural or synthetic polymeric networks. As a result of their characteristics, hydrogels develop typical firm yet elastic mechanical properties. Hydrogels have been used in biomedical applications, such as contact lenses and wound dressings. Among the advantages of hydrogels is that they are more biocompatible than hydrophobic elastomers and metals. This biocompatibility is largely due to the unique characteristics of hydrogels in that they are soft and contain water like the surrounding tissues and have relatively low frictional coefficients with respect to the surrounding tissues. Furthermore, hydrogels permit diffusion of aqueous compositions, and the solutes, there through, and have a high permeability to water and water- soluble substances, such as nutrients, metabolites and the like.

Several physical properties of the (hydro) gels are dependent upon concentration. Increase in (hydro)gel concentration may change the hydrogel pore radius, morphology, or its permeability to different molecular weight proteins. One skilled in the art will appreciate that the volume or dimensions (length, width, and thickness) of a hydrogel can be selected based on the user’s needs, such as e.g. the region or environment into which the hydrogel is to be implanted in the frame of a surgical setting. The mechanical properties of the material can be tailored according to the application site by changing the hydrogel composition (molecular chain length, crosslinking, water content and the like).

Some non-limiting examples of suitable materials constituting the microcarriers to be used in the frame of this invention include natural polymers, such as polysaccharides, co polymers of polysaccharides (cellulose, agarose, alginate, starch, chitosan and many others), polypeptides (silk, collagen, gelatin and many others), amelogenin or synthetic polymers such as polyurethanes, poly-olefins, polyethylene glycol (PEG), poly(glycolide) (PGA), poly(L-lactide) (PLA), carboxymethylcellulose (CMC) or poly(lactide-co-glycolide) (PLGA). The microcarrier may also comprise either at least one glycosaminoglycane or at least one proteoglycane, or a mixture of those two substances. The glycosaminoglycane may be for example a hyaluronic acid, chondroitinsulfate, dermatansulfate, heparansulfate, heparine or keratansulfate. The ability to use different materials is useful in different applications and adds a further degree of versatility to the device and methods described herein. In any case, the base material is not limiting as long as the other essential mechanical requirements are met, and the microcarriers can withstand a (partial) dehydration process.

Concerning the degradation/resorption rate of the carrier upon in vivo application/implant in a host, this is mainly dependent on physico-chemical properties of the polymeric material of which it is composed of, as well as further factors such as crosslinking of the polymers, the polymer concentration, the site of implant into a host and the like. The degradation/resorption rate can be calibrated by adjusting said physico chemical parameters, such as for instance by polymer crosslinking (if present), the use of inhibitor molecules, by changing the polymer density, crystallinity and/or its molecular weight distribution, changing the materials’ porosity and so forth. Generally speaking, the scaffold may be, at least in part and at least in some portion thereof, intrinsically biodegradable in vivo.

Microcarriers can exist in different shapes and porosities. In some embodiment, microcarriers according to the invention are highly porous and with an irregular shape. A preferred microcarrier in the frame of the invention comprises pores that are interconnected in order to create a continuous net of material that can act as a plausible physical support for elements such as cells or bioactive agents, while providing at the same time additional key features to the scaffold such as its softness, low resistance to interstitial flow, high compressibility, outstanding ability to regulate the capillary pressure to a constant level over a wide range of hydration states, easiness of cell/tissue invasion and so forth. The porosity of the material is preferably comprised between 50% and 99%, allowing for evacuation of liquid from the pores upon compression. Non-porous and/or regular shape microcarriers may be envisaged for use according to the invention.

For porous microcarriers, the pore size is typically comprised between lpm and 10mm, preferably between lpm and 5mm, more preferably between lpm and 2mm, even more preferably between 5pm and 500pm, obviously the size of the microcarrier itself being the upper limit of a pores size. This range of pore size is particularly convenient for a scaffold material for use in tissue engineering or regenerative medicine, since it is e.g. high enough to enable the growth of vessels through the porous material.

In the maximal hydration state of a microcarrier (that is, the volume occupied by the microcarrier submerged by a liquid with no external compression force applied thereon), the overall polymer content is comprised between 0.01% and 100% in mass of dry polymer material, preferably between 0.5% and 3%. Upon dehydration of compressible microcarriers, liquid is removed therefrom and the hydration level decreases. Microcarriers are typically used in suspension state, wherein two main regimens exist: in a dilute regime, microcarriers are separated from each others, whereas in a dense regime, a contiguous network of neighboring microcarriers is formed. The progressive transition from a dilute regime to a dense regime is called “jamming transition”, and the microcarrier concentration associated to said jamming transition can be approximately found, as will be apparent to a person skilled in the art performing a sedimentation assay. The concentration at the jamming transition state can be considered as the full hydration state of a plurality of microcarriers (100% hydration).

In the frame of the present disclosure, a “soft” material is any material that is either compressible, reversibly compressible, flexible, elastic or any combination thereof. In order to be used in living subjects, moreover, the scaffold must also imperatively be a biocompatible and/or sterilisable material suitable for medical uses. A microcarrier and/or suspension according to the invention typically behaves, depending on the hydration state, as a viscoelastic material, particularly in the dense state of suspension as defined above (100% hydration or less). Depending on the shape, porosity, density and intrinsic material properties, an elastic response can be observed for low deformations whereas a viscous behavior is observed upon high deformations. The elastic regime is important for shape stability, whereas the viscous regime is considered important for injectability and shapeability of an implant.

During or after the manufacturing process, the scaffold material can be functionalized with additional elements such as for instance bioactive molecules. Said elements can be coated on or embedded within the obtained scaffold with any suitable means known in the art, and can provide additional functional properties to the material such as enhanced/reduced biodegradation, physical stabilization, biological activity and the like. As used herein, a “bioactive molecule”, as well as “(bio)active compound” or “therapeutic agent”, is any active agent having an effect upon a living organism, tissue, or cell. The expression is used herein to refer to any compound or entity that alters, inhibits, activates, or otherwise affects biological or chemical events.

One skilled in the art will appreciate that a variety of bioactive compounds can be used depending upon the needs and circumstances. Exemplary therapeutic agents include, but are not limited to, a small molecule, a growth factor, a protein, a peptide, an enzyme, an antibody or any derivative thereof (such as e.g. multivalent antibodies, multispecific antibodies, scFvs, bivalent or trivalent scFvs, triabodies, minibodies, nanobodies, diabodies etc.), an antigen, a nucleic acid sequence (e.g., DNA or RNA), a hormone, an anti inflammatory agent, an anti-viral agent, an anti-bacterial agent, a cytokine, an oncogene, a tumor suppressor, a transmembrane receptor, a protein receptor, a serum protein, an adhesion molecule, a lypidic molecule, a neurotransmitter, a morphogenetic protein, a differentiation factor, an analgesic, organic molecules, metal particles, radioactive agents, polysaccharides, a matrix protein, and any functional fragment or derivative of the above, as well as any combinations thereof. For “functional fragment” is herein meant any portion of an active agent able to exert its physiological/pharmacological activity. For example, a functional fragment of an antibody could be an Fc region, an Fv region, a Fab/F(ab’)/F(ab’)2 region and so forth.

The bioactive compounds can be added to the microcarrier material using any suitable method known in the art, such as surface absorption, physical immobilization, e.g., using a phase change to entrap the substance in the scaffold, and the like. For example, a growth factor can be mixed with the microcarrier material composition. Alternatively, covalent coupling, e.g., using alkylating or acylating agents, is used to provide a stable, long-term presentation of a bioactive substance on the microcarrier in a defined conformation. Alternatively, non-covalent adsorbtion can be used, for example electrostatic, hydrophobic, dipole-dipole, hydrogen bonding and the like. The microcarrier are basically cell-free, but they are later on seeded in vitro or in vivo with cells after their production with the intention to transplant cells in a tissue engineering/regenerative medicine approach. Cell seeding, in its simplest implementation, involves placing the microcarriers in a cell suspension and letting the cells adhere and colonize the microcarrier. For compressible microcarriers, it is possible to greatly enhance the cell seeding process by partially dehydrating (compressing) the scaffold prior to addition of the cell suspension. In this case, the aspiration force created by the compressed microcarries tendency to expand draws the cell suspension into the microcarrier, and will seed cells deeply into the microcarrier. It is further possible to use an external perfusion device (syringe pump, pressure driven flow, and the like) to induce flow across the microcarrier and use this flow to efficiently seed cells.

In all cases, cell-adhesivity and/or porosity of the microcarrier plays a crucial role in organizing the cell seeding. Cells of a given type will only adhere in areas which are reachable for them by fluid flow, or, on a longer term, migration, and which allow for their attachment. Of note, with a porous microcarrier it is possible to sterically entrap cells even when cells do not adhere to the microcarrier material per se. Additionally or alternatively, when multiple cell types are present in a co-culture setting, one cell type can mediate the attachment of another cell type which is not normally capable of adhesion to the microcarrier.

Areas of very low pore size (comparable to cell size and smaller) will act as a complete barrier for cell seeding by flow, be it by external perfusion or by intrinsically generated flow due to microcarrier expansion. This is the case of non-porous microcarriers, where only the microcarrier surface would be colonized by seeded cells.

Different cell types or tissues fragments, e.g., stem vs. differentiated, and/or with various phenotypes in terms of differentiation, activation, metabolic or functional state, are optionally co-resident in the microcarrier of the invention. The microcarriers should be chosen to be suitable for use with any cell type or tissue fragments that one may want to transplant. Such cells include but are not limited to, various stem cell populations (embryonic stem cells differentiated into various cell types), bone marrow or adipose tissue derived adult stem cells, mesenchymal stem cells, cardiac stem cells, pancreatic stem cells, neuronal cells, glial cells, spermatozoids and ovocytes, endothelial progenitor cells, outgrowth endothelial cells, dendritic cells, hematopoietic stem cells, neural stem cells, satellite cells, side population cells. Such cells may further include but are not limited to, differentiated cell populations including osteoprogenitors and osteoblasts, chondrocytes, keratinocytes for skin, intestinal epithelial cells, smooth muscle cells, cardiac muscle cells, epithelial cells, endothelial cells, urothelial cells, fibroblasts, myoblasts, chondroblasts, osteoclasts, hepatocytes, bile duct cells, pancreatic islet cells, thyroid, parathyroid, adrenal, hypothalamic, pituitary, ovarian, testicular, salivary gland cells, adipocytes and combinations thereof. For example, smooth muscle cells and endothelial cells may be employed for muscular or tubular scaffolds, e.g., scaffolds intended as vascular, esophageal, intestinal, rectal, or ureteral scaffolds; chondrocytes may be employed in cartilaginous scaffolds; cardiac muscle cells may be employed in heart scaffolds; hepatocytes and bile duct cells may be employed in liver scaffolds; myoblasts may be used in muscle regeneration; epithelial, endothelial, fibroblast, and nerve cells may be employed in scaffolds intended to function as replacements or enhancements for any of the wide variety of tissue types that contain these cells. In general, microcarriers of the invention may comprise any cell population competent to participate in regeneration, replacement or repair of a target tissue or organ, particularly barely-reachable ones such as bone marrow, kidneys, brain, lungs or pancreas. Depending on the needs and circumstances, combinations of different kind of cells with different kind of microcarriers can be envisaged.

Microcarriers according to the invention may be prepared or chosen so that the degradation time may be controlled by using a mixture of degradable components in proportions to achieve a desired degradation rate. Alternatively, the cells themselves aid in the degradation. For example, scaffold compositions are sensitive to degradation by materials secreted by the cells themselves that are seeded within the scaffold.

The microcarriers according to the invention, as well as dense suspensions comprising said microcarriers, can be ideally transplanted on or close to a target tissue, introduced into or onto a bodily tissue/organ using a variety of methods and tools known in the art, preferably via minimally invasive surgical devices and procedures that envisage an injection step by using cannulas, syringe needles or catheters, preferably blunt ended catheters.

As used herein, the terms "patient" or “subject” refer to an animal, preferably to a mammal, even more preferably to a human, including adult and child. However, the terms can also refer to non-human animals, in particular mammals such as dogs, cats, horses, cows, pigs, sheeps, rodents and non-human primates, among others, that are in need of treatment. In some instances, a “subject” can be a plant.

As used herein, “treatment”, “treating” and the like generally refers to any act intended to ameliorate the health status of subjects such as an animal, particularly a mammal, more particularly a human, such as therapy, prevention, prophylaxis and retardation of the disease, and includes: (a) inhibiting the disease, i.e., arresting its development; or (b) relieving the disease, i.e., causing regression of the disease and/or its symptoms or conditions such as improvement or remediation of damage. In certain aspects, such term refers to the amelioration or eradication of a disease or symptoms associated with a disease. In other aspects, this term refers to minimizing the spread or worsening of the disease resulting from the administration of one or more therapeutic agents to a subject with such a disease. The term “prevention” or “preventing” relates to hampering, blocking or avoid a disease from occurring in a subject which may be, for any reason, predisposed to the disease but has not yet been diagnosed as having it for example based on familial history, health status or age.

EXAMPLARY EMBODIMENTS

The invention can best be understood from the non-limiting examples given below.

In essence, it combines a dehydration device allowing to apply controlled pressure and flow rate during the dehydration procedure, with a transfer device that can, but does not necessarily have to be a syringe like device. The dehydration device, or in the following also dehydration “box”, must be able to control final pressure and flow rate during the dehydration. In the examples below, an elementary implementation with a capillary conductor and a reservoir is given, but other solutions are also possible, for example with flow regulators driven by pressure regulators in an automated feedback loop.

In the examples below, a separate transfer device is described for transfer of the compacted microcarrier culture to an injection device such as a syringe. The transfer device may however also be directly integrated with the syringe or other in-vivo delivery device, and if cell and material losses and possible foreign particle contamination can be accepted, manual transfer would also be possible.

Example 1: Implementation of a dehydration device, transfer device and usage for injection

This example provides a basic implementation of the dehydration device, transfer tip and filling column. It also describes the dehydration process used. It makes use a specific dehydration device, which has been designed to dehydrate a solution of microcarriers at an optimal interstitial pressure for injection (and thus optimal polymer concentration). It allows an efficient removal of the dead volume while guaranteeing a safe, reproducible and easy transition from the in vitro cell culture to the in vivo injection. The whole process is harmless for the cells to be transplanted. Dehydration is complete in few minutes and the compacted microcarriers are then ready to be injected in vivo. The whole system is autoclavable (all pieces are made from polypropylene plastic) and can be treated for endotoxin removal process (with concentrated sodium hydroxide) in order to prevent excessive inflammatory response from the host.

Operating principle

Fig. 1 shows the operating principle of the drying device. The sample is applied in a drying column. A pre-set aspiration pressure is generated by a lower level of fluid in the waste reservoir of the drying box, and is transmitted to the sample via a capillary conductor (cloth). The magnitude of the aspiration pressure is set by the level of fluid in the waste reservoir, and changes little during the dehydration process since the reservoir capacity by far exceeds the liquid volume to be removed. The direction of the fluid movement is determined by the relative magnitude of the elastic expansion force of the microcarrier sample in the drying column as compared to the imposed aspiration pressure. Therefore, after equilibration, the aspiration pressure determines the fluid content of the microcarrier sample regardless of the initial fluid content. The fluid flow rate is determined by the difference of expansion pressure and hydrostatic under-pressure, but also by the total flow resistance. In this embodiment, the primary source of flow resistance is the microcarrier material near the tip of the transfer column.

Process flow

Fig. 2 provides an overview over the process flow used to obtain an injectable from microcarrier cultures. The starting material in this embodiment is a dilute suspension of carboxymethylcellulose microcarriers (CCM, synthesis given in Example 2) colonized with the cells of interest (typically, but by no means necessarily, a co-culture of HSPC and OP-9 feeder cells, Fig. 2A), but the principle is the same with any microcarrier culture (cells and carriers). This dilute suspension is transferred to the dehydration column, which consists itself of two parts, namely the loading column and the transfer tip (Fig. 2B). Excess fluid is drained into the waste reservoir by the dehydration box, until equilibrium with the aspiration pressure set by fluid level in the waste repertoire is reached (Fig. 2B). In this compacted stage, the microcarriers form a paste-like material, contained in the transfer tip. This material holds in place, such that the loading column can be removed. The transfer tip is then attached to a syringe and equipped with an injection catheter (Fig. 2C). The microcarrier-based injectable can now be transferred in vivo (Fig. 2D).

Description and usage of the drying device

This section provides further description and a detailed usage protocol for the drying device.

The drying is composed of 2 separate parts: i) a drying column and ii) a drying box (Fig. 1). The drying column handles the microcarriers, while the drying box is designed to apply a precisely known aspiration pressure and to remove excess pore fluid until equilibrium with the aspiration pressure is reached. i) Drying column

The drying columns are themselves composed of 2 parts: the loading column and the transfer tip (as shown in Fig. 2). The loading column is adapted to receive an important amount of volume, typically the content of a 6-well plate well. The transfer tip is plugged at the bottom part of the loading column. It acts as an adaptor for both the column and the needle. Once, the liquid has been removed and the microcarriers reached the desired interstitial pressure (-200 Pa for our application, corresponding to a polymer concentration of 26 mg/ml in the CCM, see Fig. 5), the particles will remain in the transfer tip. Thus, it can be unplugged from the columns and directly re-plug on a catheter or a needle.

The design of the transfer tip must be such that it maintains the compacted microcarriers when lifting the transfer tip off the capillary conductor (or filter membrane). For this, the transfer tip is designed to be conical (cone opening angles between 0° and 180°, preferentially 1° to 90°, more preferentially between 2° and 45°, even more preferentially between 3° and 30°, and most preferentially between 5° and 20°). Generally, we find larger opening angles to be necessary for larger tip opening. Alternative measures to ensure maintenance of the compacted material can be taken, such as temporary placement of a filter membrane to be removed horizontally with minimal force on the microcarrier.

Together with the capillary conductor, the tip opening is also a crucial element for regulation of the fluid flow rate. The tip opening diameter is in between about 20% of the fully expanded microcarrier size to a maximum of about 5cm, more preferentially between 50% of the average fully expanded microcarrier size and 1cm, and even more preferentially between 100% of the average fully expanded microcarrier size and 0.5cm. ii) Drying box

The box is composed of a drying platform and a waste reservoir. The capillary conductor (contacting cloth) ensures the connection between these two compartments. The capillary conductor is optionally also active as a filter to maintain the particles in the transfer tip during dehydration. In this case, its pore size show be adapted to exclude particles from entering the capillary conductor; depending on particle elasticity, it is sufficient to have a pore size smaller than the microcarrier size, or it is necessary to have it much smaller. For elastic porous particles of a mean size of about 0.5mm, we find a mean pore size of about 0.1mm to be sufficient. Alternatively, a filter membrane is imposed between capillary conductor and transfer tip; in this case, the minimal requirement on the capillary conductor is merely not to lose all of its pore water at the desired set pressure to maintain sufficient hydrolic conductivity for the dehydration process. The filter membrane in that case should have a pore size at most on the size scale of the microcarriers, preferably much smaller (including down to nanometric sizes such as standard 0.22 micrometers or 0.45 micrometer filter membranes).

The waste reservoir is a Falcon tube into which a whole at a given height has been drilled to ensure a constant level of saline buffer whatever the exact amount of excess medium in the CCM solution. The excess of liquid will indeed leak out of the tube by the hole performed at a specific height defined by the experimental needs. Indeed, polymer concentration may vary from one application to another. We use here a drying box with a preset height of hole at 2cm equivalent to about 200 Pa aspiration pressure), which we find to produce a CCM concentrate suitable for a subcutaneous injection while preserving cell viability.

Usage protocol:

We use the following protocol to concentrate the CCMs into an injectable paste under sterile conditions:

1) Under sterile conditions, humidify the capillary conductor with saline buffer. Add saline buffer in the waste reservoir in order to reach the hole level.

2) Plug the transfer tip to the loading column. Then, place the drying column in the holder. Make sure that the column is in contact with the capillary conductor.

3) Using a serological pipette, transfer the content of the well in the loading column and avoid formation of air bubble in the column. Otherwise, tap gently on the column in order to remove them. 4) Dehydration should take few minutes. In the meantime, prepare the injection syringe (typically 1 ml syringe) by aspirating sequentially 150 ul of non-seeded CCMs at 26 mg/ml and 150 ul of air.

5) Once the CCM reached the desired concentration, using sterile forceps, unplug the transfer tip from the loading column. Specific care should be taken to avoid CCM fall off from the tip.

6) Plug the transfer tip containing the CCMs to the catheter or the needle. Plug the injection syringe.

7) Insert the catheter into the insertion hole (the injection site should be cleaned and insertion hole should be made before the dehydration).

8) Inject the content of the syringe.

9) Remove the catheter gently.

Fabrication details

This section provides technical details and protocols on the custom fabrication of the loading columns and the drying box.

Loading columns

Fig. 3 shows the technical dimensions of a drying column, which consists of two mandatory pieces: the loading column and the transfer tip. Optionally, it can be closed with a cap on its top.

In the following, a laboratory-scale fabrication method for the loading column and transfer tip are given.

Loading columns:

Materials

• Falcon 15 ml (bottom and top part) (PP)

• 1000 ul tip from Axygen (7.9cm) (PP)

Fabrication method

1) Cut the falcon 15 ml in 2 parts: bottom part at the 3 ml line and top part at the 12 ml line. Discard the middle part.

2) Assemble the 2 parts by melting a rim of plastic with a hot air gun, followed by rapidly pressing the two parts together.

3) Cut the very bottom part (conic part) of the falcon tube. The diameter of the hole should be the same than the top part of the 1000 ml tip

4) Assemble the tip and the falcon tube, again by melting a rim of plastic on both the tip and the shortened Falcon tube with a hot air gun followed by pressing.

5) Cut the bottom part of the tip (remove between 1.2-1.5 cm). Outer diameter 0.4 cm. Average total height size of the loading column 11.5 cm

Materials

• 20 ul from MultiGuard, Sorenson, Bioscience Inc. (PP) Fabrication method

1) Cut the top part at 1.2 cm from the top (largest part)

2) Then cut the bottom part at 2.3 cm from the bottom (the narrowest part), below the filter.

Drying box:

Materials

• 1000 ul pipet box, Axygen scientific (PP). The tips are not needed.

• Table cleaning cloth, local supermarket (Coop, 82%viscose, 18% polypropylene, autoclavable)

• 50 mL falcon tube, Corning (PP)

Fabrication method

1) Make a hole in the tip shelf of the size of a 50 ml falcon tube

2) Insert the falcon tube and melt it with a soldering iron in order to attach it to the box

3) Make a hole in the falcon tube at the desired height (here 2 cm) from the top part. This will define the aspiration pressure

4) Contacting cloth: cut in a table cleaning cloth a rectangle of 8x7 and with a tail of at least 6 cm at one side.

Aspiration pressure and polymer concentration

The use of the drying device hinges on the equilibrium relation between polymer concentration and aspiration pressure. To establish this relation for the biomaterial constituted by the dense suspension of CCM, we measured the dry polymer content at different preset aspiration pressures. We did so for uncoated, cell-free CCM precursor material.

For this measurement, we replaced the transfer tip by a cell strainer. In this way, we could determine the weight of material equilibrated with a predefined series of aspiration pressures, before extensive deionized water washing to remove free salts, and microwave or oven drying to constant weight for dry weight determination.

Fig. 5 shows the relationship between the dry weight polymer concentration of the biomaterial and the aspiration pressure as set by the height difference Ah between the shelf of the pipet box and the lower edge of the pressure regulator hole. At a preset Ah of 2cm, we obtain a polymer concentration of 26 +/- 3mg/mL. This is about the double of the total polymerizable mass content of the premix used to make the bulk scaffolds before fragmentation, indicating that there is substantial compaction and probably compression of the CCMs. However, this is still substantially below the about 40mg/mL of polymer concentration where an onset of decrease in viability was seen (Example 3). In any case, using the partial dehydration procedure as described here, we did not observe a decrease in cell viability

Addition of viscosants

In some applications, and in particular to maintain the fluid associated with the microcarriers during extrusion or injection, it is advantageous to render the binder fluid more viscous, for example by adding methylcellulose, carboxymethylcellulose, agarose, starch, polyethylene glycol or any of a large number of cell compatible vicosing agents. Typically, one wishes to increase the viscosity of the interstitial fluid by a factor of 2x to lOOO’OOOx in such applications. For the lower viscosities, addition to cell culture medium may be sufficient. For higher viscosities (lOx, and especially lOOx increase or more, it is advantageous to add the viscosant only towards the end of the dehydration procedure, or dehydrate the microcarriers to a slightly higher pressure (for example l.l-5x) than finally desired and use the rehydration induced by lowering the dehydration pressure for rehydration with the viscosant (as one would for rehydration with any other fluid).

Bibliography example 1

1 Moosbrugger, C. in Atlast of Stress-Strain Curves (ed C. Moosbrugger) (ASM International: The Materials Information Society, 2002).

2 Beduer, A. et al. A compressible scaffold for minimally invasive delivery of large intact neuronal networks. Adv Healthc Mater 4, 301-312, doi: 10.1002/adhm.201400250 (2015).

3 Beduer, A., Verheyen, C. A., Bonini, F., Burch, P. & Braschler, T. submitted (2019).

Example 2: Synthesis of compressible microcarriers

The aim of this example is to estimate the admissible linear compression in sample microcarrier system for co-culture of hematopoietic stem cells and the stromal OP-9 line.

Microcarrier synthesis

Compressible carboxymethylcellulose scaffolds in accordance with W02017/029633 are produced by cryogel bulk scaffold synthesis. For this, a reaction mix consisting of 13.56mg/mL carboxymethylcellulose (AQUALON CMC 7LF PH, 90.5 KDa, DS: 0.84) and 0.486mg/mL adipic acid dihydrazide, buffered with 6.3mg/mL PIPES neutralized to pH 6.7 by 1.2mg/mL NaOH was prepared and filtered through a 0.22um filter (Stericup). After activation by 2.7mg/mL l-Ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDC), the mix was frozen at -20°C in 30mL syringes. After 48h of ciyo-incubation, the syringes are thawed. The scaffolds thus obtained are then fragmented to irregular, compressible microcarriers by extrusion through a 22G catheter. This yields a suspension of microscaffolds suitable for use as microcarriers. These microcarriers are then extensively washed, and autoclaved for sterilization.

To allow stromal cell (OP9) adhesion, the surface of the sterile microcarriers is covalently modified with collagen type 1 (from bovine skin, Sigma, C4243), producing collagen- coated carboxymethylcellulose microcarriers (CCM). For this, we used a previously published protocol with adaption to the particulate nature of the CCMs (Serex et al., 2018)(Xia et al., 2017). Briefly, we dehydrated the microscaffolds using a cell strainer in a SteriCup filtration system (C3240). They were washed sequentially with D1 water and acetic acid buffer (pH=4, 100 mM). Then, the microscaffolds were immersed in a coating solution containing 10% collagen type 1 (mass of protein/dry mass of microscaffolds) diluted in acetic acid buffer (pH=4, lOmM). After coating, the collagen-coated carboxymethylcellulose microscaffolds (CCM) were rinsed twice with D1 water in order to remove the excess of non-adsorbed proteins. Thereafter, covalent crosslinking of the collagen was performed by immersing the CCMs in a solution containing EDC (1 mg/ml) and MES buffer (pH 4.5, 100 mM) in D1 water for 10 minutes. Finally, the CCMs were abundantly rinsed with D1 water and a solution of Na2C03 (pH 11, 100 mM) and kept in PBS at 4°C.

Example 3: Microcarrier culture and viability

Microcarrier culture

The aim of this example is then to evaluate stability of the collagen-coated carboxymethylcellulose microscaffolds CCM as described in Example 2, seeded with cocultures of murine stromal OP9 and hematopoietic stem and progenitor cells (murine hematopoietic stem and progenitor cells or HSPC, obtained by sorting bone marrow extract for ckit⁺, lin-, Sca-1⁺ “KLS”). Seeded at an initial OP9:KLS ratio of 100:1, the CCMs were cultivated for 3 months to achieve steady-state cellular composition prior to mechanical and cellular stability testing against controlled uniaxial compression. This data allows to follow the fate of the co-cultures on the CCMs during controlled compression, and thus to estimate the allowable compression during injection.

For this, collagen-coated CCMs were seeded using 75,000 OP9 (GFP) and 7,500 KLS (DsRed) per mg of dry scaffold weight. After initial incubation to allow for cell adherence, the scaffolds were distributed in ultra-low adhesion plates; a 6-well plate (2mg of dry scaffold/well) and a 24 well plate (0.5mg of dry scaffold per well). Culture was performed by adding first 3mL respectively 0.5mL of conditioned medium, and then another 3mL respectively 0.5mL at day 7, followed by half-media changes every week for 3 months. We took care to avoid aspiration of the CCMs by holding the plate in a slanted position for about 30s prior to medium aspiration. This allows the CCMs to sediment and be protected from aspiration. The procedure however is expected to remove a sizeable fraction of the cellular descendance generated by in-vitro hematopoiesis at each medium change.

Scaffolds were retrieved at 3 months of culture and pipetted into an observation chamber adapted for removal of excess liquid during the concentration process (Fig. 6A, exploded view).

Setup for evaluation of viability

This chamber consisted of a laser cut Perspex sheet in the shape of a microscope slide (i.e. 75mm x 25mm), with a rectangular central observation area (30mm x 12.5mm), closed with a floor consisting of a glued coverslide (Fig. 1A). In addition, we placed a laser-cut cleaning cloth made from polypropylene and cellulose (Migros, Switzerland) into the central area (Fig. 6A). The cleaning cloth matched the dimensions of the central observation area (nominally 30mm x 12.5mm, de facto about 0.25mm smaller at each edge due to the finite laser spot size), but in addition we cut a circular hole at its center (8mm nominal diameter, real diameter estimated about 8.5mm). This circular hole serves as sample concentrator, but also as holder for uniaxial compression with a 8mm circular chuck (Fig. 6B). After compression, the samples can be observed by confocal microscope (Fig. 6C). Prior to scaffold placement, we hydrated the cleaning cloth with the circular hole with cell culture medium (i.e. 50/50 mix of conditioned and fresh medium as described in the main text) supplemented with 50mM HEPES for use in the absence of a CO2 enriched atmosphere. We then slowly pipetted the suspension of CCMs from the cultures into the central hole. To avoid compression during transfer, we aseptically broke open a medium aspiration pipet for this purpose. At the same time, we removed excess medium by aspiration from the cleaning cloth. Taking care to always leave sufficient medium to have the gap between the observation chamber wall and the cleaning cloth filled with medium, we avoided excessive capillary pressure on the accumulating material. In the rudimentary setup used here, without proper pressure control, this step proved quite difficult, indicating the need for a more precise way of dehydration to reproducibly achieve a desired living biomaterial consistency.

We obtain an assembled biomaterial where the particles have about their natural size, as needed to estimate approximately the permissible compression. By sedimentation of CCMs (unseeded) in large excess of medium, we estimated the polymer concentration at full expansion to be about 8mg/mL (see below, section “Uniaxial compression properties”). We added enough material to completely, but not excessively fill the sample hole with cell-seeded biomaterial; we repeated the experiment 2x, once with material from the 6-well culture, once from the 24-well culture.

Fig. 7 shows the detailed setup used for uniaxial sample compression. After mounting of a 8mm cylindrical chuck onto a TextureAnalyzer XTPlus machine, the machine is calibrated for force gain by the standard calibration routine in the Texture Exponent user interface. A microscope coverslide identical to the one used for compression chamber fabrication is then placed onto the sample stage of the TextureAnalyzer XTPlus machine, and chuck height calibration is carried out. In this way, the zero-height is defined at upper surface of the coverslide. Once the calibration routines successfully carried out, the compression chamber with previously loaded sample is placed under the chuck, giving rise to the compression setup as shown in Fig. 7. The chuck is then lowered to establish contact with the sample, taking care to avoid air bubbles. Then, the chuck is moved vertically until zero net force is detected; this is done by a built-in routine for this purpose available in the TextureExponent program.

From this zero-force equilibrium height, uniaxial compression is carried out. Three heights are tested: 0% compression, corresponding to no further movement of the chuck; 75% compression, corresponding to a decrease of the sample height to 25% of the original sample height above the coverslip surface; and 100% compression, corresponding to theoretical lowering of the chuck to the coverslip surface. In practice, even at 100% theoretical compression, some unknown, but minimal sample height is maintained due to mechanical compliance in the TextureAnalyzerXT Plus machine. The 0% percent control is to assess potential damage done to the cells by the contact with the chuck. All compression is carried out a speed of O.Olmm/s, to emulate quasi-static conditions.

Mechanical data acquisition and treatment Force-distance data was acquired during the compression phase. This was done purposefully at relatively high data rate (200 samples per second). We could then lower the noise to a level compatible with straightforward curve analysis by applying a Gaussian low-pass filter (with a width, as expressed by the standard deviation parameter, of 40micrometers or equivalently 800 measurements).

Viability and cellular composition

For each of the two experiments done, we assessed viability and cellular composition initially (after chamber loading), at 0% compression (chuck zeroing), at 75% compression and at 100% compression. This implies that we followed 2 samples through the entire process of loading, chuck zeroing, and compression to 75% and then to 100%.

For viability estimation, we used both extrinsic staining by the nuclear stain Hoechst 328 and loss of intrinsic fluorescence (GFP for OP9, DsRed for KLS and descendance). We had indeed noticed on preliminary trials (100% compression vs. samples directly from culture) that Hoechst 328 strongly stains the nuclei of dead cells, and weakly are the ones of live cells and the scaffold material. Upon cell death, on the other hand, the GFP signal is lost rapidly, followed on the time scale of about 20 minutes by loss of DsRed as well.

Hence, we could obtain an estimate of the dead cell fraction as well as an indication of the composition (OP9 vs. KLS and descendance) of the remaining live cell population under different conditions by counting predominantly red, predominantly green, and predominantly blue cells on the confocal images.

In practice, we acquired 5 to 7 confocal stacks with at least 8 slices (Zeiss LSM700 microscope, Plan-Apochromat 20x with numerical aperture of 0.8, area of 640microns x 640microns, z-spacing 9.25 microns) for each sample and the 4 different conditions: A) sample loading only; B) sample loading plus chuck positioning; C) 75% compression; D) 100% compression. We repeated the experiment twice, once with a sample assembled from CCMs grown in a 6-well plate, once with a sample from the 24-well culture.

The observation chambers were kept closed with a second coverslide during transport and microscopic observation; between experiments, medium was replenished if necessary by placing a drop of medium (50/50 mix of conditioned and fresh medium with 50mM HEPES as described above) such as to keep the clefts between cleaning cloth and Perspex sheet fully filled, indicating minimal capillary pressure. This was necessary since opening and closing by sliding the closing coverslide (Fig. 1C) removed a small amount of medium sticking to the coverslide.

Cell population at 3 months

From the confocal images acquired before any contact with the compression chuck (Fig. 8 for an example), we could estimate the cellular population adherent to the scaffolds at 3 months. On average, we find an approximately equal amount of red (44%+/-7%, HSPC) and green cells (49%+/-8%, OP9) and a smaller fraction of dead cells (7%+/-3%). The result suggests that on the long run, the OP9 can sustain about a 1:1 loading with firmly attached HSPC (t-test of the proportion of green vs. red cells per stack, with Moulton correction for clustering in two experiments: P=0.28, which is compatible with the null hypothesis of a 1:1 association and therefore equality of the two proportions).

We also find many cells floating freely in the wells, but due to the regular half-media changes and also the removal of most of the medium during reconstitution of a semi-solid biomaterial in the sample area of the compression chamber, we expect the quantification on the scaffolds to include nearly only cells that are firmly attached at 3 months.

Of note, in addition to well-identifiable round DsRed cells, we also find a variable proportion of DsRed that seems to be distributed in small vesicles in the cytoplasm of the corresponding OP9 cell. Although we did not investigate the phenomenon any further, this could be endo-lysosomal processing of dead DsRed positive cells.

Cell viability during compression

Fig. 9 shows confocal images illustrating the effect of various degress of uniaxial compression (0%, 75%, 100%) on the co-cultures of OP9 (green) and KLS+ descendance (red). Fig. 9A and Fig. 9B show that qualitatively, mere contact with the chuck (Fig. 9A) and even transient uniaxial compression by 75% of the original sample height (Fig. 9B) has only minor effects on the co-cultures, whereas 100% compression kills a large fraction of the cells.

Fig. 10 shows a quantitative analysis of the percentage of dead cells compared to the estimated total number of cells. While no significant difference can be detected between the material simply loaded into the observation chamber (“Control: Loading” in Fig. 10) and contact with the compression chuck but no actual compression. We detect a significant increase in the dead cell fraction after the 75% uniaxial compression, although the co-culture still remain largely intact (raise of the dead cell fraction from 7% to 13%). Theoretically complete compression on the other hand leads to the immediate death of more than 50% of the cells.

Composition of the viable cell fraction during compression

Beyond cell viability, we also assessed the composition of the remaining viable cell population in terms of hematopoietic cells vs. OP-9 stromal cells. It could indeed conceivable be that compression, while maintaining viability of the attached cells, preferentially expels one cell type or the other. To guard against this possibility, we evaluated the fraction of red fluorescent DsRed+ cells among the total viable cell fraction at different levels of compression, and also after loading of the chamber only.

Fig. 11 shows the results regarding the composition of the remaining viable cell fraction. This fraction is composed of distinct green (GFP+, OP9) and red (DsRed+, KLS and descendance) fluorescent cells. As outlined above, at 3 months of culture, there is a nearly 1:1 ratio of red and green cells (“Control: Loading”). We find a slight, but statistically significant decrease of the DsRed+ fraction after the chuck zero-force equilibration procedure (from 53% to 45% on average, P-value of 0.04 associated with equilibiration vs. initial loading in linear regression with both the culture and equilibration vs. initial loading as explanatory variables, Bonferroni correction for a total of three tests against the 0% compression condition). Compression by itself then has little influence on the average DsRed+ fraction, although the variability increases especially for the 100% compression, where cells are killed in entire image areas.

From the data presented in Fig. 11, we conclude that there is no major bias in the composition of the viable cell fraction upon compression. This result has to be interpreted within the context of a culture with regular half-media change, and dehydration during loading into the observation chamber. Indeed, we expect most of the free-floating or loosely attached cellular descendance of the hematopoietic stem and progenitor cells to be lost during culture, and at latest during chamber loading. The cells which are present after chamber loading are therefore relatively firmly adherent cells. Compression of the scaffolds with this firmly adherent cell populations seems to kill the cells located in unfavorable areas, rather than to preferentially affect one cell type or the other.

Uniaxial compression properties

The uniaxial compression experiments directly yield force-distance graphs: while the chuck slowly compresses the sample, the TextureAnalyzerXTPlus machine measures the force the sample opposes to compression.

After application of a low-pass filter to smooth the noisy, but high-bandwidth force measurements, we converted the force-distance curves to stress-strain diagrams. The stress s is indeed calculated by relating the measured force Fto the cross-sectional area of the chuck A (Moosbrugger, 2002): s = - F . eq. 1

A

The strain e in turn is obtained by relating the compressive chuck displacement Ah to the sample height ho :

Eq. 1 and eq. 2 correspond to the so-called « engineering » (Moosbrugger, 2002) stress and strain, since the original sample dimensions are used in their calculation(Moosbrugger, 2002).

Fig. 12 shows the stress-strain diagrams for the four compression experiments: to 75%, and then again from original sample height to 100% compression, for the two samples prepared (one from the 6-well culture, one from the 24- well culture).

All four compression experiments show a qualitatively similar behavior in compression: with increasing strain, the elastic recoil force raises more and more steeply. Quantitatively, the compression curves show high repeatability within each sample, as the compression to 100% recapitulates closely the previous compression to 75% on the common part from 0 to 75% compressive strain. This shows the very good elastic recovery typical for cryogels. There is a more important difference between the first and second sample, the second appearing stiffer. This is most likely rooted in a difference in particle concentration achieved during loading. This phase is indeed manually controlled by the effective dehydration pressure applied, which is only controlled to within visual observation, avoiding high capillary pressure as evident by the formation of strongly curved menisci in the clefts between cleaning cloth and Perspex sheet.

At least during the initial stages of uniaxial compression, we expect mostly pore fluid to be expelled from the highly porous biomaterial. This estimate the effective polymer concentration as a function of the strain as defined by eq. 2: eq. 3 where Co is the polymer concentration in a reference state. We carry out the loading procedure such as to minimize capillary aspiration pressure by careful observation of the menisci. Hence, we expect a material density close to one observed during spontaneous sedimentation, which by dry mass determination is about co=7mg/mL crosslinked material in cell culture medium.

Noting that the concentration of polymer changes due to progressive compression allows to estimate the variation of the Young modulus at each polymer concentration from the stress-strain curves shown in Fig. 12. The Young modulus is defined as the change of stress per unit of change of strain: da eq. 4 de

Fig. 13 outlines the Young moduli as estimated from eq. 4 plotted against the polymer concentration as estimated by eq. 3. Over a wide range, a 3^rd power law is observed for both samples, in agreement with a similar law reported for bulk ciyogel scaffolds. (Beduer et al., 2015a) The polymer concentrations shown on the x-axis of Fig. 13 should be considered indicative: It is quite difficult to determine the exact polymer concentration reached when loading the polymer chamber. For the small volumes used for the compression measurement, the mere act of transferring to a scale causes pore fluid to be lost. We performed two experiments to define the upper and lower bounds of the polymer concentration after loading.

To estimate the lower bound, we performed sedimentation over 24h of a dilute suspension of CCMs in a large excess of cell culture medium. From the volume of the sediment and the determination of the dry mass present (by the CCMs with excess deionized water followed by oven drying), we could estimate the free sedimentation concentration to be 6.0+/-0.6mg/mL.

To estimate an upper bound, we replicated the loading of the compression chamber with larger volumes (ca. lmL) and a 40 micrometer cell strainer together with a larger mock compression chamber to be able to transfer with less change of fluid volume. The cell strainer makes the transfer to a scale for hydrated weight determination easier, although some loss of fluid is still expected. In this way, we could estimate an upper bound to the chamber loading concentration to be 9.6 +/-2.0mg/mL.

The actual polymer concentration is expected to between the upper and lower bounds thus determined. As a reasonable estimate, we indicate the range midpoint and the upper and lower bounds as extremes, and therefore assume the loading concentration to be 8+/- 2mg/mL.

Using this estimation of 8+/-2mg/mL for the loading polymer concentration, Fig. 13 indicates that the implants have Young moduli in the lower kPa range (1.2+/-0. 6kPa). Also, there is well-respected 3^rd power law for the dependency of Young modulus on the polymer concentration throughout a large part of the polymer concentration range throughout the compression experiments. We previously found a 3^rd for bulk ciyogel scaffolds(Beduer et al., 2015a). The 3^rd power reflects the mechanics of a porous elastic structure(Beduer et al., 2015a) where increasing compression leads to loss of pore fluid without major change of the mechanical structure. Important deviation from the 3^rd power law only sets in above about 100-200mg/mL; this concentration corresponds about to previously reported polymer concentration within the ciyogel walls of bulk scaffolds(Beduer et al., 2015a), and thus departure from the 3^rd power-law can be taken as a sign of complete pore closure. This is in line with the viability data: at 75% compression (corresponding to about 40mg/mL polymer concentration), most of the pores conserve some pore fluid, so that the cells remain protected(Beduer et al., 2015a). Pore closure and ensuing massive cell death only sets in at higher polymer concentrations still.

For in-vivo injections, we concentrate the CCM suspensions by applying an aspiration pressure of about 2 cm of water column (about 200Pa). During this procedure, we reach a polymer concentration of about 26mg/mL (Example 2). This is well within the domain of the 3^rd power in Fig. 13, and also below the 32mg/mL associated with 75% compression where much of the viability was still conserved.

Overall, we can conclude the cells should remain protected from excessive compression in the biomaterial formulation used for in-vivo injections because most of the pores remain at least partially open, so that little mechanical force is transmitted to the cells.

Example 4: Hematopoietic stem cell transplantation

Aim

Modeling the interaction between the supportive stroma and the hematopoietic stem and progenitor cells (HSPC) is of high interest in the regeneration of the bone marrow niche in blood disorders. In this example, we present an injectable co-culture system to study this interaction in a coherent in vitro culture and in vivo transplantation model. We assemble a 3D hematopoietic niche in vitro by co-culture of supportive OP9 mesenchymal cells and HSPCs in porous, chemically defined collagen-coated carboxymethylcellulose microscaffolds (CCM, Example 2). Flow cytometry and hematopoietic colony forming assays demonstrate the stromal supportive capacity for in vitro hematopoiesis in the absence of exogenous cytokines. After in vitro culture, we recover a paste-like living injectable niche biomaterial from CCM co-cultures by controlled, partial dehydration. Cell viability and the association between stroma and HSPCs are maintained in this process. After subcutaneous injection of this living artificial niche in vivo, we find maintenance of stromal and hematopoietic populations over 12 weeks in immunodeficient mice. Indeed, vascularization is enhanced in the presence of HSPCs. Our approach provides a minimalistic, scalable, biomimetic in vitro model of hematopoiesis in a microcarrier format that preserves the HSPC progenitor function, while being injectable in-vivo without disrupting the cell-cell interactions established in vitro.

Scaffold fabrication

We used here the microcarriers of example 2. Collagen CCMs surface coating We coated here the microcarriers of example 2. Animals

All experimental procedures were approved by the Animal Care and Use Committee of the Canton of Vaud (ACUC, Vaud, Switzerland). All animals were hosted in the EPFL facilities and were kept under a controlled 12 hours light/dark cycle and at constant room temperature 22+/-2°C. DsRed C57BL/6JRj (DsRed) adult male mice were sacrificed and tibiae and femurs were collected for DsRed+ HSPC isolation. 8-16 weeks old NOD SClD-y (NSG 5557, Jackson laboratories) immunodeficient female mice (n = 3 control; n = 6 experimental) were used as recipients in the transplantation model.

Culture of OP9 stromal cells

Using an established murine mesenchymal stromal cell line (OP9s) (Nakano etal., 1994), cells were expanded at 70-80% confluency for one to two weeks in alpha-minimum essential media (a-MEM) plus Glutamax (32561, ThermoFisher), 10% fetal bovine serum (FBS, 10270-106, G1BCO), and 1% Penicillin/Streptavidin (P/S, 15140122, Thermo Fisher Scientific). OP9 cells were donated from the Daley laboratory (McKinney-Freeman et al., 2009), who received them directly from the Nakano laboratory (Nakano et al., 1994). They were transfected at passage 7-10 with a constitutively expressed GFP lentiviral construct, and expanded. Cells were kept at 37°C and 5% CO2, and were passaged every 2-4 days at 3:1 or 4:1 ratio, until they reached 70-80% confluency. OP9s were not kept in culture for more than three weeks before use in experiments. Cells were washed with lx PBS (10010056, Life Technologies) and tiypsinized with 0.05% Tiypsin-EDTA (25300054 Life technologies), counted, and kept in suspension on ice prior to use.

Isolation of HSPCs DsRed adult mice were euthanized with CO2 according to approved protocols and both tibiae, femurs, and pelvis extracted. After cleaning the bones of all soft tissue, they were kept in PBS on ice until all bones were isolated. Bones from age and gender-matched C57BL/6J wild type controls were isolated in parallel for fluorescence-activated cell sorting (FACS) single-color controls. Bones were subsequently crushed using a mortar and pestle in buffer solution (PBS, ImM EDTA (15575020, Thermo Fisher Scientific), 2% FBS), until no large chunks of cells were visible. All cell isolation steps were carried out on ice. Cells in suspension and the crushed bones were washed through a 70pm cell strainer and spun down (10 min, 300g, 4°C). The cell pellets were resuspended in red blood cell lysis buffer (420301, BioLegend) for 30 seconds, before being diluted with buffer solution, and spun down again (5 min, 300g, 4°C). The cell suspension was stained with a lineage antibody cocktail, washed, and incubated with magnetic beads according to manufacturer’s instructions for hematopoietic Lineage depletion (Lin depletion kit : 558451; BD Pharmingen). The total bone marrow cell pellet was resuspended in 3mL volume and loaded into a magnetic separation cell Sorter (AutoMACS, Miltenyi) to remove all lineage positive (Lin+) cells in suspension. The resulting cells were then blocked for 15 minutes on ice (5pg/ml hlgG; I4506-10MG, Sigma Aldrich), and finally stained for one hour on ice with lineage Streptavidin-PO (1/200), as a conjugate to label any remaining Lin+ cells, as well as c-Kit PE-Cy7 (1/200), Sca-1 APC (1/100). After washing the stained cells with buffer solution and straining through a 85pm filter, the cell suspension was run through a FACS system (Aria Fusion) and the resulting Lin-, c-Kit+, and Sca-1+ (KLS) cells were sorted into Iscove’s Modified Dulbecco’s Medium (IMDM) + Glutamax, 25mM HEPES (31980022, Life Technologies) supplemented with 10% FBS and 1% P/S. In total, 2-3 adult male DsRed+ mice (aged 8-12 weeks) were euthanized to collect approximately 200,000 KLS+ cells in suspension for each experiment. After FACS, cells were kept on ice for approximately 1-2 hours until co-seeding with OP9s on the scaffold.

Co-culture of HSPCs and stromal cells

All cells for co-seeding experiments were cultured in 50% fresh basal media (1MDM + Glutamax 25mM HEPES, 10% FBS, 1% P/S) and 50% conditioned 1MDM media (CM). Conditioned media was obtained by culturing confluent GFP+ OP9s with IMDM media for two days (48 hours), filtering the conditioned medium, and freezing the media for no longer than two months at -20°C. After HSPCs and OP9s were collected in suspension and counted, cells were kept on ice for maximum 1 hour.

3D co-seeding: Collagen-coated microscaffolds (CCMs, 13.5 mg/ml in PBS) were dried using a cell strainer in a SteriCup filtration system (C3240), using an autoclave cloth to transmit the capillary pressure. Once dried, the globule of CCMs was transferred to a 6- well ultra-low adhesion plate (Corning, CLS347) using the tip of a 2 mL stereological pipette. For each condition, the two cell types (HSPCs, OP9s) were combined, spun down, and re-suspended in a minimal amount of media (approximately 100 pL in total). For 2 mg of scaffold per well (dry weight), the following cell ratios were used for the 1:10 “high” seeding (150,000 OP9; 15,000 HSPCs) and 1:100 “low" seeding (150,000 OP9; 1,500 HSPCs). After adding the cell suspension (~100 uL) to the dried scaffold on the ultra-low adhesion plates, the CCMs with cells were incubated for 1 hour at 37°C and 5% CO2. After 1 hour, 3 mL of 1MDM media (50% conditioned, 50% fresh) was added per well. Co seeded CCMs were then left in culture for 12 days, with a supplementary dose of 3 mL 1MDM media (50% conditioned: 50% fresh basal media) at D7, without removing any of the previous media. of 120,000 cells per well in a 6-well plate 1 hour prior to HSPC seeding (~ 13,000 cells/cm²), using 1MDM media (50% conditioned: 50% fresh). HSPCs were seeded at the previously established co-seeding ratios, 1:10 “high” (12,000 HSPCs per well) and 1:100 “low” (1,200 HSPCs per well). If limited by HSPC cell number, the 2D condition was performed with only the low seeding density. Cells were fed at D7, complementary to the 3D culture timeline, with 3mL added and no media removed, then cultured at 37°C and 5% CO2 for 12 days in total. Co-seeding experiments were repeated in at least two separate experiments, with technical triplicates within each experiment.

Compression testing was performed to assess the limit of compression compatible with cell survival in example 3.

Isolation of cells at D12

Cells were collected for each condition (high/low; 2D/3D) in three fractions: cells in suspension, cells adherent to the CCMs, and any cells adherent to the ultra-low adhesion plates. For cells in suspension, media was collected in a 50 mL falcon tube, and cells were washed and collected twice with serum-free media. For the CCMs adherent fraction, cells were detached via enzymatic digestion as follows. CCMs were transferred to 24 well plates with a 1000 um pipette tip and 1 mL of collagenase 1 (17100-017, ThermoFisher Scientific) 0.04% was added per well of CCMs for 25 minutes at 37°C and 5% CO2. The collagenase digestion was stopped with media complemented with serum, and a stereological pipette was used to dissociate any cells from the CCMs. Finally, cells in solution were run through a 100 pm cell strainer and collected for further manipulation. Trypsin digestion was used for the very limited number of cells adherent to the ultra-low adhesion plates. Cells were spun down and re-suspended in a buffer solution. At this point, each of the conditions (4 total - 2D/3D, High/Low Seeding Density) and each replicate (n = 3 per experiment; two independent experiments) were processed for flow cytometry and for methylcellulose colony forming unit (CFU) assays. Each fraction (non-adherent suspension, collagenase-digested CCM-associated, and bottom-adherent trypsin- digested) was processed separately and the results are presented for all fractions compounded after analysis, based on the total number of cells recovered per well for each of the fractions. Flow cytometry

Cell suspensions for each condition and fraction were re-suspended with blocking solution (5pg/ml hlgG) for 10 minutes at room temperature and then Lineage cocktail was added (1:20 dilution) for 20 minutes on ice. After washing with buffer solution and filtered with a 85pm cell strainer, cells were pelleted and re-suspended with antibody mix for 45 minutes on ice. The antibody mix contained: c-Kit (PE-Cy7; 1:200), Sca-1 (BV711; 1:50), CD45 (AF700; 1:100), CD3 (BV421; 1:50), B220 (PE-Cy5; 1:50), CDllb (APC-eFl. 780; 1:1000), and Grl (APC; 1:500), diluted in BD Brilliant Stain Buffer (563794, BD Pharmingen). The antibody mix also contained a 1:4 dilution of BrightCount beads (Invitrogen, C36950). Cells were then diluted with DAP1 solution (PBS-EDTA and DAP1- UV at 1:5000) and run through a BD LSR 11 SORP flow cytometer, while resting on ice during the stain preparation.

Methylcellulose CFU Assays

Single cell suspensions, as isolated in 3D (non-adherent, scaffold-adherent, and bottom- adherent) and 2D (non-adherent and bottom-adherent), were separately kept on ice. Each fraction was plated in 1.1ml methylcellulose (M3434, STEMCELL Technologies) in duplicate for each condition and each replicate for hematopoietic clony forming untit /CFU) assay (Mcniece et al., 1990). CD45+ cells were counted with FACS using BrightCount beads (Invitrogen, C36950), to be able to back-calculate the exact number of cells plated. At Days 7 and 10, each CFU plate was read using a StemVision instrument (Stem Cell Technologies), and total colonies were assessed automatically (StemVision proprietary software) and verified manually on the acquired high-resolution whole-plate images according to colony number, size, and cell distribution (Mcniece et al., 1990).

Subcutaneous Transplantation

After 14 days of in vitro culture, seeded-CCMs in suspension were collected from the well plate and poured into the column of the drying device (Example 1) allowing for the CCMs to settle down into the reservoir and reach the desired interstitial aspiration pressure (ca. 200Pa) and therefore polymer concentration (26 +/- 3 mg/ml, Example 3). This condenses the CCMs into a paste-like material with a Young modulus of 1.2+/-0.6 kPa [Example 3 ), which we find sufficient to sustain a 3D architecture in vivo. Sterile syringes were used to aspirate 0.1 ml of coated scaffold without cells, followed by 0.1 ml of air to ensure separation between them. The reservoir with the sedimented cultured scaffold (50 pL) was connected to a lmL syringe and a 20G flexible catheter (BD Biosciences 381703) was plugged in the other end.

NSG mice were chosen for the experiments as OP9 stromal cells are derived from a mixed genetic background and therefore purely syngeneic transplantation was not possible. Prior to injections, anesthesia was induced in NSG mice with 4% isoflurane USP- PPC (Animalcare Ltd). An ophthalmic liquid gel (Viscotears, Alcon) was used to protect the eyes and local isoflurane was reduced to 2%. Mice were placed on a heating pad to keep the temperature constant during intervention, and the back of each mouse was shaved at the area of the injections. Betadine (Mundipharma Medical Company) was spread onto the shaved regions to disinfect the skin.

To perform the subcutaneous injection, a small orifice was created in the disinfected skin using a 18G needle and the 20G catheter (Tro-Vensite i.v. canula, Troge, Hamburg), connected to the loaded syringe, which was gently inserted subcutaneously about 2 cm from the pierced skin. For each mouse, two separate injections of 50 pL each were performed subcutaneously on either side of the spine. No sutures were required. At the end of the procedure, the mice were placed back in the cage grouped per condition. The entire preparation of the scaffolds and all the injections were performed under the hood to ensure sterility throughout the whole procedure. Each injection, from start to finish, lasted less than 20 minutes per mouse.

Animals were treated with antibiotics in drinking water consisting of 30 mg of Enrofloxacin (300 pL of Baytril 10% ad us. vet, 100 mg/mL, Bayer) and 5 mg of Amoxicillin (100 pL of Amoxi-Mepha 200mg/4mL, Mepha Pharma AG) as well as 500 mg of Paracetamol (Dafalgan®) in a total of 250 mL sterile water for the entire duration of the study and replaced every 7 days. Animals were monitored daily by the researchers, and after two weeks, they were monitored daily by animal care services.

Sacrifice and samples harvesting

NSG immunodeficientmice were euthanized 12 weeks post-injections through inhalation of CO2 (6 minutes). The back was shaved gently to better localize the two implants. The samples were harvested in each mouse being careful to keep some subcutaneous tissue around to study the integration of the scaffolds within the normal tissue. Samples were then fixed for 24 hours in 4% paraformaldehyde at 4°C (10 mL PFA in 15 mL Falcon tube), washed three times with PBS, and embedded in paraffin.

Histology

Tissues were fixed in paraformaldehyde (PFA), submitted for stepwise dehydration and embedded in paraffin blocks for sectioning at 3-4 pm thickness with a rotary microtome (RM, Leica microsystems) . After floating on a water bath to flatten, sections were mounted on glass slides (Superfrost+ slides, Menzel glaser). Paraffin sections were stained with Hematoxylin and eosine (H&E) using the Tissue-Tek Prisma automate (Sakura) and permanently mounted using the Tissue-Tek glas G2-coverslipper (Sakura) to assess morphology. Detection of rabbit anti-GFP (Abeam, ab6673, diluted 1:400), rabbit anti- Dsred (MBL, PM005, diluted 1:500), rat anti-CD31 (clone SZ31, Dianova, DIA-310-M, diluted 1:50), rabbit anti-vWF (Abeam, ab9378, diluted 1:100) or rabbit anti-Perilipin (Abeam, ab3526, diluted 1:200) was performed manually. After quenching with 3% H202 in PBS lx for 10 minutes, a heat pretreatment using 0.1M Tri-Na citrate pH6 was applied at 60°C in a water bath overnight. Primary antibodies were incubated overnight at 4°C. After incubation of a goat, a rat or a rabbit Immpress HRP (Ready to use, Vector Laboratories), revelation was performed with DAB (3,3'-Diaminobenzidine, Sigma- Aldrich). Sections were counterstained with Harris hematoxylin and permanently mounted. Number of CD31+ stained vessels were counted in 2-7 images (500x500pm) covering the entire area of the sectioned scaffold (amounting to 2-7 images per scaffold depending on the size of the scaffold) for four scaffolds each in both the unseeded and seeded conditions.

Microscopy and image quantification

Co-seeded CCMs were kept in 3D culture for 12 days in vitro. At days 1, 4, 7, and 11, a small volume of suspended CCMs were removed and transferred to a deep cavity glass microscope slide (produced in lab for imaging, see Example 3, Fig. 6). Within an hour after transferring CCMs to the imaging chamber, they were imaged at varying magnifications (20X for imaging quantification; 63X for cell morphology) using a Zeiss LSM 700 Inverted Confocal Microscope. As the cells were endogenously labeled for GFP (OP9s) and DsRed (HSPCs), serial imaging was conducted at each time point without significant cellular manipulation. Each image acquired was kept to the same microscope settings as the DO condition, though as more cells proliferated on the CCMs, laser power was changed to allow for clear image acquisition. In image processing, the ratio of DsRed+ to GFP+ cells was accounted for in order to compare cell proliferation over time. At end point, the seeded CCMs were stained with Hoechst 328 (0.1 pg/ml) to visualize the scaffold filaments themselves.

Images used for quantification were composed of a 25-z-stacked, volume- rendered image. To analyze the data, each fluorescent channel was separated and the compiled volume-rendered image was used. Each image/channel was analyzed using Fiji/lmageJ’s threshold tool, with the resulting quantified fluorescent areas converted to cell numbers by using the mean area per cell, as established by manual identification of a subset of the cells. For each 3D experiment, a total of eight independent CCMs were analyzed from two experiments to plot the relative cell proliferation over time.

Statistical Analysis

Values are shown as mean plus or minus the standard deviation (mean +/- SD). Student’s t-test was performed for all experiments when comparing two conditions only, or a Two- Way AN OVA for multiple conditions over time. The p-value for statistical significance was p<0.05.

RESULTS: The study aimed to provide a microcarrier co-culture system for convenient and minimally invasive injection of a tissue-like living biomaterial, without disrupting cellular viability and multi-cellular interactions during the injection procedure. We simultaneously seeded stromal OP9s and HSPCs on porous CCM microscaffolds (Fig. 14A). The system self-organized such that the OP9 stroma lined the scaffolds coated with collagen 1 to support the HSPC subpopulations (Fig. 14B), and allowed for in vitro studies in diluted microcarrier suspension cultures. For subsequent validation in vivo, the intact co-cultured CCMs, together with their cellular payload, were dehydrated by a custom dehydration device (Fig. 14C, Example 1) and delivered in vivo by subcutaneous syringe- injection (Fig. 14D).

Experimentally, our starting biomaterials are highly elastic and porous microscaffolds consisting of crosslinked carboxymethylcellulose. A 3D view based on confocal reconstruction after staining with rhodamine 6G is provided in Fig. 14E. The microscaffolds are designed to be reversibly compressible. This allows for facile exchange of the pore fluid by arbitrary sequences of dehydration and rehydration. To obtain the collagen-coated microscaffolds CCM, we made use of such cycles to efficiently and covalently functionalize the microscaffolds with collagen type 1 to provide native stromal cell adhesion motives.

For the purpose of scaffold seeding, we generated green fluorescent protein (GFP+) OP9 cells and obtained red fluorescent primary murine HSPCs (cKit+Lineage- Scal+, referred to as KLS) from the marrow of DsRed C57BL/6JRj mice. We then co seeded a mixture of OP9 stromal cells and KLS+ cells into our CCM scaffolds by making use of the spontaneous aspiration capacity of dehydrated scaffolds to distribute the cells throughout the microscaffolds. In vitro culture of this system demonstrated self organization into a stromal compartment, adopting the scaffold architecture and hosting the HSPCs and their progeny (Fig. 14F, Fig. 14G).

Collagen-coated, mesenchymal stromal cell-seeded scaffolds promote hematopoietic cell proliferation over time

We first assessed the capacity of the artificial stroma, consisting of CCMs lined with OP9 marrow stromal cells, to support hematopoiesis in vitro. We made use of the constitutively expressed fluorescent proteins to allow for the visualization and semi- quantitative analysis of co-cultures by confocal microscopy (Fig. 15).

Indeed, confocal imaging showed effective spreading of OP9 stromal cells on the CCM scaffolds and attachment of the co-seeded HSPCs to the OP9s, with continuous proliferation of hematopoietic cells within the scaffold over 11 days in culture (Fig. 15A). The stromal cells are essential, since at 24 hours HSPCs failed to adhere to the CCMs in the absence of the OP9 cells (Fig. 15B, Fig. 15C). In the presence of OP9, we found distinct signs of HSPC supportive characteristics within our in vitro hematopoietic niche analog. This included HSPC nestling within OP9 stromal cells (Fig. 15D) and colony formation within the matrix after 3-4 days of culture (Fig. 15E). Together with the absolute requirement for stromal cells in the absence of exogenous cytokines, this indicated intimate and favorable 3D interactions between the two cell types when loaded into the CCMs.

We then quantified the relative proliferation of the HSPCs in our system. To do so, we seeded the CCMs at two relative HSPC densities: high (1:10) and low (1:100) seeding density (HSPCs to OP9 cells). We followed the relative proliferation of the HSPCs and their progeny by evaluating the area occupied by red fluorescence (DsRed) as compared to green fluorescence (green) in confocal sections of the live co-seeded CCMs over time. By also measuring the average area occupied per HSPC and OP9 (180 pm² +/- 90 pm² for the HSPC vs. 1270 pm² +/- 460 pm² for the OP9, n = 8 cells in each case), we converted the area measurements to number ratios of HSPC:OP9. Fig. 15F shows that the ratio of HSPCs (and their descendant progenitors) to OP9 cells within the CCMs increased from 0.07+/- 0.02 to 1.3+/-0.4 for the high seeding density (intended initial 0.1=1:10 ratio) during 11 days of culture. Likewise, the number ratio HSPCs (and their descendant progenitors) to OP9 increased from 0.018+/-0.01 to 0.12+/-0.03 for the low seeding density (1:100).

This indicated that the seeding density after the initial adhesion step, followed by dilution of the CCMs into a large volume of medium, reflected the original 1:10 and 1:100 ratios. In addition, at both seeding densities, a significant increase in relative numbers of HSPCs and their progeny took place within the CCMs, consistent with effective multiplication and expansion of the HSPCs and their differentiated progeny supported by the OP9 cells in an exogenous cytokine-free environment. However, this microscopic assay underestimated the HSPC multiplication, as numerous cells were observed outside of the CCMs in suspension. Moreover, it does not account for the proliferation of OP9- BMSCs. Due to the constitutive expression of DsRed, it is uninformative regarding the fate of the HSPCs in regards to differentiation or maintenance of stem cell properties, which was thus evaluated through surface protein phenotyping and functional colony assays.

Functional quantification of hematopoietic proliferation on stromal-cell seeded scaffolds as compared to 2D cultures

We next aimed to better characterize the fate of the HSPCs during in vitro culture. For this purpose, we used flow cytometry to analyze the co-cultures in 3D on CCMs, and compared them to corresponding standard 2D controls with the same 1:10 and 1:100 initial HSPC:OP9 seeding densities. The constitutive expression of GFP and DsRed by the OP9 cells and HSPCs/progeny, respectively, allowed for the visualization and separation of the OP9 stromal versus the HSPC-derived hematopoietic populations. For example, the dot plot in Fig. 16A shows the composition in terms of GFP and DsRed expressing cells for a 3D, 1:10 experiment after 12 days of culture. We found that the HSPCs and their progeny, as identified by the DsRed⁺GFP- population, consisted nearly exclusively of CD45+ cells, as expected for all cells of the hematopoietic lineage derived from bone marrow HSPCs (Fig. 16B) (Weissman and Shizuru, 2008). These cells, which expressed no lineage markers in the cell surface at seeding (Linage- is part of the KLS definition), consisted at day 12 on a mixture of lineage negative and lineage positive cells (Fig. 16B). Acquisition of major lineage markers thus revealed hematopoietic differentiation within the CCM coculture. Preservation of c-kit+ expression in a subset of the lineage negative DsRed⁺GFP- CD45+ cells (Fig. 16C) suggested the preservation of progenitor function, as later tested in Colony-forming unit (CFU) assays.

Quantitative comparison showed that the total CD45+ cell expansion was significantly higher in 2D 1:10 “high” (254.0±164.4) compared to 3D 1:10 “high” (32.0±13.0, p=0.0094) cultures, as well as in 2D 1:100 “low” (530.0±126.3) versus 3D 1:100 “low” (19.0±11.3, p<0.0001) cultures (Fig. 16D). Very interestingly, although fold expansion of total CD45+ cells was consistently higher in 2D versus 3D conditions, fold expansion of the hematopoietic progenitor compartment as reflected by quantification of phenotypic progenitors (CD45⁺Lin cKit⁺ population) was similar for 2D (4.1±3.0) and 3D (2.6±1.2) cultures of the 1:10 “high” seeding density (Fig. 16E). Progenitor expansion was higher in 2D versus 3D in the 1:100 condition (4.1±2.7 versus 0.4±0.3, p=0.0092). However, total cell recovery from the 3D microscaffolds was challenging to achieve, and thus quantification of hematopoietic expansion may have been underestimated for the 3D conditions, especially for the scarce CD45⁺LinxKit⁺ progenitors in the 1:100 condition, which tightly associate to the stroma to form cobblestone-like colonies.

In order to complete the analysis with functional hematopoietic potency in the co cultures, we performed CFU assays, which quantify the number of oligopotent progenitors able to form functional hematopoietic clonal colonies after a 7-10 day culture in semi-solid, cytokine-rich media. Like for the FACS analysis, 2D and 3D HSPC/OP9 co cultures seeded at “low” (1:100) and “high” (1:10) relative HSPC:OP9 ratios were enzymatically digested to obtain a single cell suspension after a 12 day co-culture and plated for CFU assay. Fig. 16F shows an example of a CFU assay at 7 days. For the purpose of comparison and to obtain the total number of CFUs produced by each culture condition, CFU counts were normalized per 1000 HSPCs seeded with OP9s at Day 0 (1000 KLS = starting HSPC concentration, n = 6 for each condition). The number of CFUs was not significantly different (p=0.166 via Two-Way ANOVA) between the 2D and 3D conditions at each seeding density (1:10 or 1:100, Fig. 16G), which is congruent with the CD45+ckit+ hematopoietic progenitor quantification by flow cytometry for the 1:10 “high” density (Fig. 16B). The number of relative colonies is significantly higher in the “low” (1:100) seeding condition (2D: 1093.86 ± 567.88, 3D: 764.69 ± 306.84) as compared to the “high” (1:10) condition (2D: 408.34 ± 126.32, 3D: 369.42 ± 99.46), suggestive of either nutrient competition or a negative paracrine regulation by differentiating hematopoietic cells at higher CFU seeding densities. Compared to controls obtained by direct plating of fresh KLS cells without culture prior to the methylcellulose assay, we observed a maintenance, or minor expansion, of the functional HSPC compartment. This is expected and reflects a biomimetic, homeostatic condition, as there is no exogenous addition of any of the cytokines commonly used for in vitro HSPC expansion (e.g. thrombopoietin, stem cell factor, or Fms-related tyrosine kinase 3 ligand, (Costa et al., 2018)). The aim here was indeed to provide a minimalistic system enabling further screening without interference from exogeneous cytokines. Overall, we conclude from the in vitro functional characterization of our HSPC/OP9 co-cultures that 3D co-culture in CCMs promotes similar expansion of functional hematopoietic progenitors to conventional 2D conditions, with a reduced output of differentiated CD45+Ckit- hematopoietic cells. Injection of co-seeded CCMs demonstrates both OP9 and donor hematopoiesis in the engineered niche over 12 weeks in vivo in NSG immunodeficient mice

Compressible porous scaffolds have previously demonstrated promise in providing an injectable solution to traditional cell-based tissue engineering techniques (Beduer etal., 2015b; Bencherif et al., 2012b). In addition to the preservation of hematopoietic progenitor function in the 3D HSPC/OP9 co-culture conditions, we were interested in testing the biocompatibility and hematopoietic support capacity of the HSPC/OP9-loaded CCMs in vivo.

Fig. 17A-D shows the workflow for transitioning from a microcarrier-like suspension culture to a transplantable co-culture biomaterial. After a predefined time of in vitro culture as a dilute suspension (Fig. 17A), the CCMs are collected and dehydrated to a controlled level by a device specifically designed for that purpose (Fig. 17B, and detailed information in Example 1). The device applies a pre-set hydrostatic pressure to the CCMs by means of a capillary conductor. As the hydrostatic pressure sustained by the biomaterial constituted by the CCM is strongly linked to its concentration, this ensures constant material consistency compatible with regards to injection and hematopoietic niche reconstitution in vivo. Here, we set a hydrostatic pressure difference of 0.2kPa, equivalent to a fluid level difference of about 2cm, to concentrate the co-culture biomaterial to 26 +/- 3 mg/mL (Example 1). At this concentration, the material remains easily injectable and matches the kPa range for the vascular part of the bone marrow niche (Bello et al., 2018), as detailed in Example 1.

After concentration of the co-culture biomaterial, the transfer tip is fitted onto a syringe, and assembled with a catheter (Fig. 17C) for subcutaneous injection (Fig. 17D). By performing the injection into a cell-culture dish containing medium, we assessed whether the procedure of partial dehydration and passage through the catheter during the injection would be harmful to the cells (Fig. 17E). We found that we are able to retain high viability of cells immediately after injection and 24 hours-post injection, as compared to the pre-injection controls in both the GFP+ OP9s and DsRed+ HSPCs (Fig. 17E; p = 0.822 via Two-Way AN OVA).

For assessment of the in vivo performance of the co-culture biomaterial, we injected the living co-culture biomaterial prepared via the dehydration device into the subcutaneous dorsal region of NSG mice. Supplement 2 includes a video of the entire dehydration and injection procedure. The implants were easily visible externally up to the end of the 12-week follow-up period (Fig. 17F). At sacrifice, skin flaps from the back of the animal revealed macroscopic vascularization of the injected scaffolds within the dermal tissue of animals (Fig. 17G).

We carried out two independent sets of experiments to assess the in vivo evolution of the co-culture biomaterial. In both sets, the samples consisted of CCMs seeded with HSPCs and OP9s, at the 1:10 initial seeding ratio and cultured for 14 days prior to partial dehydration and implantation. In the first set, we included a control group of scaffolds seeded with OP9 only, whereas in the second set, the control group consisted of completely cell-free CCMs. The aim was to dissect the effect that the scaffold alone, the OP9-seeded scaffolds, or the OP9/HSPC co-seeded scaffolds would have on the in vivo outcome. In both sets, after 12 weeks in vivo, hematoxylin and eosin (H&E) standard histological staining revealed intact scaffold particles (Fig. 18A, D and G). In all conditions, the scaffold was host to diverse cell types. Immunohistochemistry (IHC) against GFP revealed strong persistence of confluent OP9 stroma across the scaffolds shown by the large areas stained in brown (Fig. 18E and H). Such staining was absent from the initially cell-free CCM implants (Fig. 18B), providing evidence for the specificity of the anti-GFP staining. Similarly, anti-DsRed IHC revealed a positive signal only for scaffolds loaded with both GFP+ OP9s and DsRed+ HSPCs (Fig. 181), indicating persistence of hematopoietic cells from the donor DsRed+ HSPCs 12 weeks after implantation. Lack of DsRed+ signal on scaffolds loaded with only OP9 cells (Figure F) and initially cell-free implants (Fig. 18C) attests to the specificity of the anti-DsRed IHC.

The HSPC/OP9-loaded scaffolds macroscopically appeared to vascularize better and conserved a higher proportion of donor-derived stroma (Fig. 19F and I) than cell-free scaffolds (Fig. 19C). Indeed, HSPC/OP9-seeded scaffolds showed a 6.6 times higher vascularization (p=0.003, t-test with Welch approximation after log transform) than cell- free scaffolds, as quantified by relative number of CD31+ vessel segments (Fig. 19J).

In the study groups, we observed multinucleated DsRed+ cells, suggesting the presence of specialized donor-derived hematopoietic progeny within the implants, morphologically reminiscent of megakaryocytes, the specialized sessile hematopoietic cells responsible for platelet production. In order to assess megakaiyocytic differentiation within the implant, we therefore performed von Willebrand factor (vWF) IHC stains (Fig 6A, D and G). Only HSPC/OP9-seeded scaffolds contained highly positive vWF+ cells (Fig. 19D and G), indicative of megakaryocyte lineage commitment within the scaffold, and thus of active in situ hematopoiesis. Finally, in order to assess adipocytic differentiation within the implanted scaffolds, we performed IHC stains with perilipin (Fig. 19B, E and H). HSPC/OP9-seeded scaffolds (Fig. 19E and H) but not the unseeded controls (Fig. 19B), presented frequent areas of adipocyte ghosts with a cytoplasmic perilipin positive signal characteristic of mature adipocytes. In summary, we can conclude from in vivo experiments that murine HSPC/OP9-seeded CCM scaffolds can be implanted in NSG mice to produce highly vascularized structures which retain donor stroma and contain locally active hematopoiesis as well as interspersed adipocytes, features reminiscent of adult marrow (Weiss, 2008).

DISCUSSION:

In this study, we developed an easy-to-use system enabling smooth transition from in vitro co-culture to an injectable thatself-assembles in situ to recapitulate structural and mechanical features of the hematopoietic marrow. Its aim was to bridge the gap between various defined co-culture systems in vitro and more realistic but complex in vivo niches such as long-described heterotopic ossicle formation (Tavassoli and Crosby, 1968; Maniatis etal., 1971; Friedenstein etal., 1982). In orthopedics, porous scaffolds have long been used in conjunction with BM aspirates to enhance bone formation (Yoshii et al., 2009). More recently, they have also been identified as a solution enabling localization of BM niches to a biomaterial, enabling thorough in vitro testing of scaffolds prior to implantation, and also targeted implantation (Shah et al., 2019; Shih et al., 2017). Yet, due to the bulk format of these scaffolds, it is difficult to perform live imaging or high- throughput screening on them. We therefore sought to combine the advantages of a microcarrier culture system with the mechanical robustness of a bulk scaffold for hematopoietic niche engineering.

To achieve this goal, we based our system on compressible sub-millimetric carboxymethylcellulose scaffolds. In vitro, they can be used as classical microcarrier cultures. Additionally, in vivo, these specifically engineered CCM scaffolds self-assemble by microscaffold interlocking. The process creates a mechanically stable implant, inducing colonization with a fibrovascular stromal tissue. This dual behavior enables smooth injection of arbitrary scaffold volumes, followed by in situ regeneration of a porous niche structurally resembling trabecular bone, with mechanical moduli in the vicinity of lkPa (measured ex-vivo, Example 3), as reported for the vascular niche of the bone marrow (Bello et al., 2018). From a practical point of view, CCMs are easily fabricated with standard equipment (freezer, autoclave for sterilization, laminar flow hood for coating under sterile condition), such that production is easily scalable at affordable costand compatible with Good Manufacturing Practice (GMP) production.

To enable the adhesion of the stromal OP9 cells, the CCMs feature covalently bound collagen I (Beduer et al., 2018; Serex et al., 2018). Indeed, among a series of extracellular matrix molecules, collagen type I has been shown to provide the highest proliferation levels with KLS cells (Choi and Harley, 2017). Further, contrary to Matrigel used for generation of hematopoietic ossicles (Bello et al., 2018; Hughes et al., 2010; Reinisch et al., 2017), collagen I is a single protein of defined composition that is amenable to clinical use (Lecarpentier et al., 2018; Salvade et al., 2007). OP9 cells have been shown to direct pluripotentstem cells towards the hematopoietic fate, and also have an ability to maintain engraftable hematopoietic stem cells in in vitro 2D co-cultures systems for up to 2 weeks in the absence of additional cytokines (Naveiras, 2009). In this study, the combination of OP9 cells and CCMs was found not only to enable the baseline culture of HSPCs over 12 days in culture, but also to provide for easy and effective implantation of the hematopoietic scaffolds in vivo for a follow-up of 12 weeks in NSG mice.

By stable transfection of OP9 with GFP and the usage of KLS from DsRed mice, the CCMs were readily imaged in 3D culture, and serial imaging gave qualitative insight into early-stage HSPC nestling and cobblestone-like colony formation within the 3D scaffolds. We believe this is indicative of a similar hematopoietic supportive behavior from the stromal cells as compared to 2D controls. We further analyzed the cellular populations phenotypically by flow cytometry, and functionally by CFU assays. We found the output of the 3D cultures, measured as total fold hematopoietic expansion (CD45+), to be lower in 3D than in 2D (Fig. 16D). However, the HSPC compartment, as measured by the CFU counts in the 2D versus 3D cultures (Fig. 16G), is essentially unaffected by this lower output of total hematopoiesis. As the 3D cultures are effectively concentrated in a much smaller volume with higher local cytokine levels (Rodling et al., 2017), our results point towards differential regulation of hematopoiesis in our configuration leading to HSPC enrichment, which may be ascribed to the difference balance of the hypothesized “vascular” versus “endosteal” niches (Leisten etal., 2012; Sanchez-Aguilera and Mendez- Ferrer, 2017).

Our CCM-based co-culture system was not only amenable to live, high-resolution imaging on sub-samples obtained by simple pipetting, but also allowed for standardized aggregation prior to injection by the use of a specifically designed dehydration device. This avoided loss of cell viability during preparation and injection. At the end of the experiment, the scaffold aggregates were both visible beneath the surface of the mouse’s skin and were also easily identifiable with integrated host vasculature after opening of the dermal tissue (Fig. 17). Histological staining showed the presence of the GFP+ OP9 with colonies of the DsRed+ HSPC interspersed throughout the recovered tissue sections. Moreover, these implanted CCMs demonstrated no morphologically visible innate inflammatory foreign body response or fibrous capsule surrounding the injected materials. This unique finding supports the hypothesis that homeostatic extramedullary hematopoiesis may be engineered in vivo in the form of subcutaneous implants, in the absence of ossification surrounding a marrow cavity, analogous to the soft-tissue masses of benign extramedullary hematopoiesis, often intermixed with adipose tissue, that occur in humans upon extreme hematopoietic demand (Roberts et al., 2016). Previous systems have employed calcified bone surrounding an engineered niche to model hematopoiesis in vitro or to encourage hematopoiesis once implanted in vivo (Torisawa et al., 2014; Holzapfel etal., 2015; Shafiee etal., 2017; Ventura Ferreira et al., 2016; Blache etal., 2016; Shih etal., 2017; Reinisch etal., 2017; Bourgine etal., 2018). To our knowledge, the system presented here is the first of its kind to show hematopoiesis after long-term in vivo subcutaneous transplantation of BMSCs without any calcified bone component.

In this study, we were also cognizant of the multi-cellular complexities associated with BM function, along with recent models incorporating BM adipose tissue in vitro (Henriksson et al., 2017; Fairfield et al., 2019). We found mature adipocytes within the scaffolds transplanted with both OP9s and HSPCs, indicating that our model may be of use to study the complex relationships between hematopoiesis and adipocytes in vitro and in vivo (Mattiucci et al., 2018), a field that has been hampered by the difficulty of immobilizing mature adipocytes in co-culture.

Future work follows the premise of further recapitulating the BM microenvironment by manipulating culture conditions, including addition of hematopoietic cytokines, and the modification of the stromal compartment by replacing the OP9 in the CCMs with a human CAR cell line equivalent for a minimalistic niche, or possibly a mix of cell lines to better reflect the niche heterogeneity. Moreover, by using primary human samples, we may be able to further address the limitations of patient- derived xenograft PDX models (de la Puente et al., 2015; Sanchez-Aguilera and Mendez- Ferrer, 2017; Shah and Singh, 2017; Song et al., 2019). In this respect, our system offers new perspectives for personalized-medicine techniques with the possibility for high- throughput screening in vitro followed by validation of selected treatments in a direct implantation in vivo. CONCLUSION:

In summary, we present in this report a novel 3D co-culture system of HSPCs and BMSCs for studying BM hematopoiesis on chemically defined, collagen-coated cryogel- based scaffold microcarriers. The method for co-seeding two cell populations of the BM is simple and scalable, requiring no exogenous cytokine supplementation for hematopoietic progenitor cell maintenance and proliferation. We further designed a dehydration device enabling on-the-fly preparation of a paste-like injectable implant from dilute suspension cultures. Through minimally invasive subcutaneous transplantation of this living implant, both the stromal and hematopoietic cell populations were able to survive in vivo for 12 weeks, showing incorporation into the native tissue via de novo vascularization and positive staining for donor GFP+ (OP9 BMSCs) and donor DsRed+ cells (HSPC progeny), including megakaryocytes. Moreover, our tissue engineered BM has provided a first indicator of supporting subcutaneous, extramedullary hematopoiesis in healthy adult murine tissue without simultaneous ossification. We observe some induction of adipogenesis, pointing towards bidirectional communication between the niche and the hematopoietic compartment. A tool allowing to study such bidirectional signaling could be invaluable in both research on PDX models and radiation-induced bone marrow adiposity. In conclusion, this 3D engineering BM niche demonstrates promise to better model the BM microenvironment through a defined in vitro to in vivo transition, enabling future work in fundamental and patient-specific applications in hematology and bone marrow biology at large.

In this example, we compare injectability of cytodex3 vs. CCM (synthesis see Example 2) microcarrier biomaterials in an in-vitro assay. Some injectability is preserved with both materials, showing the generic nature of the compacting process, and its applicability to different microcarrier materials. The CCM material however does perform better in protecting cells, showing that in addition to the general process, the nature of the microcarrier also has some influence (porous microcarriers are better than non-porous, flexible microcarriers better than rigid ones).

The performance of the CCM with a co-culture of OP9 and hematopoietic stem and progenitors cells can be seen in Fig. 17E: dehydration and ejection have no impact on cell coverage of the carriers, indicating perfect protection (the DsRed and GFP labeling of the two cell types are rapidly lost upon cell death).

The performance of the cytodex3 dehydrated material is slightly less good (Fig. 20): As for the CCM material, the dehydration device performs perfectly, but a significant amount of cells (OP9) are lost during injection. This is truly cell loss, as the amount of cell death on the cytodex beads does not increase (Fig. 20). Even so, the amount of cells conserved as measured by the GFP area confluence remains significantly above 0, demonstrating that the general method of dehydration and injection can clearly still be used with cytodex 3 microcarriers. Example 6: Cell survival in-vivo in different materials

Fig. 21 compares relative in-vivo transfer efficiency for different materials. From Fig. 20, we anticipated some cell loss for microcarriers that are not specifically adapted for the purpose and therefore seeded lOx more cells than on the CCM carriers (Example 2).

The preliminary results shown in Fig. 21 indicate that there is about an order of magnitude of cell loss for the cultisphere G, cytodex 3 and also a non-porous CCM analog. Nevertheless, sufficient cells survive to deliver a living implant, indicating applicability of the method to different microcarriers even though it is evident that engineering appropriate properties into the microcarriers themselves increases the efficiency of the process. On the CCM carriers, OP9 show long-term engraftment in NSG mice (3 months; see also Fig. 18).

Example 7: Flow rate characterization during dehydration

This example provides characterization of the dehydration box, transfer tip and drying column as described in Example 1, for dehydration of CCM microcarriers as synthesized in Example 2. These data show that for a wide range of set target aspiration pressures (indicated as cm H20 where 1cm H20 is about 98 Pa), the volumetric flow rate is independent on the applied pressure (Fig. 22A). For low microcarrier concentration as typical for dilute suspension culture, there is also relatively little variation of the flow rate with varying polymer concentration, although the flow rate will decrease at higher polymer concentration when approaching the equilibrium.

Example 8: Layered compacted microcarrier culture.

Commercial BC1 were expanded in NeuroBasal/DMEM F12 culture medium supplemented with B27 and N2 complement, lOng/ml FGF2 and EGF and pen/strep. BC1 cells were harvested using Accutase and seeded on collagen-functionalized microcarriers (Figure AA). This experiment shows that the BC1 do not adhere onto the selective microcarriers, indeed some die and others form clusters. Figure AB shows the layered compacted microcarrier culture. For this human foreskin fibroblasts (HFF) were cultured in DMEM F12 supplemented with 10% FBS and then harvested using trypsin. These cells were first seeded on the collagen-functionalized microcarriers. The otherwise non adherent BC1 were then seeded onto the living microcarrier in order to form the layered compacted microcarrier culture shown in Figure AB.

Example 9: Integrity of the microcarrier system after exogenous or endogenous reloading

This example provides evidence for the integrity of the microcarrier system, once injected, to accept reloading at a second timepoint, whether by reinjection or by colonization of local resident cells that acquire the capacity to colonize the microcarrier through the presence of the stromal layer.

For Fig 25 A-C a total of 0.4 ml of the partially dehydrated CCM microcarrier suspension was delivered subcutaneously as described extensively in examples 3 and 4. After the microcarrier system had acquired its definitive geometry, a suspension of 40 microliters of cell-emulating low diffusion carbon microparticles was carefully injected (Fig. 25A). As shown in Fig 25B, the microcarrier system preserved its geometry after receiving the secondary graft. Fig 25c shows how the cell-emulating microparticles delivered at a second timepoint could distribute through the previously injected microcarrier while preserving the microscopic architecture.

For Fig. 25D-E 30-50 microliters of the partially dehydrated CCM microcarrier suspension were injected intra-bone, through the murine tibial plate, into the bone marrow cavity. Fig. 25 D-E show at low and high magnification the microcarrier suspension integrating into the bone marrow hematopoietic tissue and thus the potential of the so-delivered microcarrier suspension to incorporate the functional progenitor populations in situ.

Luhmes organoids were commercially obtained from Neurix SA, Geneva, Switzerland. They were layered below microcarriers functionalized with Matrigel and then subjected to partial dehydration and culture at a negative pressure of -0.9kPa on a diffusive membrane, while providing fresh medium at a flow rate of 100 micrometers/s on the other side of the membrane. These culture conditions were maintained for 7 days, before fixing the samples with 4% PFA, and processing for paraffine embedding. The samples were stained for Bill tubulin and DAP1. The results of the experiment (Fig. 24 C) show nearly complete integration of the microcarriers into the organoids (80-90% integrated area compared to original size of organoid) with maintenance of organoid viability. Other control experiments without negative pressure or only minimal negative pressure (insert culture pressure, 0 to -O.lkPa), little integration occurred despite the microcarrier functionalization (<20%). This shows the utility of the present invention to obtain microcarrier-tissue composites.

Description of the Figures

Fig. 1: Schematic representation of the hydrostatic pressure applied by the device. A capillary conductor drives the pressure applied by the level in the waste reservoir of the drying box to the sample in the drying column. The pressure can easily be adapted by changing the level of liquid in the waste reservoir.

Fig. 2: Overview of the transition from in vitro culture to in vivo injection. A) In vitro co culture of OP9 (stromal cells) and HSPC (KLS) inside our collagen-coated CCM. The biomaterial is used as a microcarrier in a diluted form. B and C) Concentration the diluted CCM by the drying device. The protocol is detailed below. D) Subcutaneous injection of the concentrated CCM in mice.

Fig. 3: Technical details of the drying column. The drying column consists of a loading column and a transfer tip. The transfer tip is reversible attached to the loading column by pressing. Optionally, the loading column can be closed by a cap. Fig. 4: Technical layout of the drying box. The drying box a customized pipet tip box: a waste reservoir with a lateral hole for pressure regulation is inserted into tip shelf, and a capillary conductor in the form of specifically cut wiping cloth is laid out onto the plateau. An extended part of the capillary conductor extends into the reservoir to transmit the aspiration pressure set by the position of the pressure regulator hole. The pipet box also serves as a containment for the excess liquid leaving the waste reservoir by the pressure regulator hole.

Fig. 5: Relation between aspiration pressure and the dry weight polymer content of the CCM biomaterial (uncoated precursor). The aspiration pressure is expressed by the height difference between the pressure regulator hole and the plateau of the drying box (Ah in Fig. 4).

Fig. 6. Observation chamber adapted for uniaxial compression. A) Fabrication: The chamber is assembled from a cleaning cloth, a Perspex sheet and a microscope coverslide. The Perspex sheet is cut to the outer dimensions of a microscope slide (75mm x 25mm) with a C02 laser cutter, with a central observation area (30x12.5mm). It is closed on the lower side with a microscope coverslide, which is glued to the Perspex sheet with silicone glue. The assembly is completed with a cleaning cloth, cut to fit into the observation area, and with a central sample area of circular shape (8mm) diameter. B) Use in compression. Using an 8mm diameter chuck on a texture Analyzer XTPlus machine, sample in the sample area can be compressed uniaxially. Excess liquid is taken up by the cleaning cloth. C) After compression, the sample can be observed by confocal microscopy. For this purpose, the chamber can be closed reversibly by a second microscope coverslide to limit evaporation.

Fig. 7. Uniaxial sample compression. A sample is loaded into the circular sample area (Fig. 6, here in cross-section), defined within the cleaning cloth. The compression chamber is then placed onto the sample stage of a mechanical testing machine (TextureAnalyzer XTPlus) and then compressed by uniaxial movement of the chuck. Excess liquid squeezed out from the pore space during compression is drained by the cleaning cloth.

Fig. 8. Z-projection (maximum intensity) of a confocal stack of a co-culture of OP9 and KLS on CCMs at 3 months. Hoechst 328 was used to stain for nuclei of dead cells.

Fig. 9. Z-projection (maximum intensity) of a confocal stack of co-cultures of OP9 and KLS on CCMs after uniaxial compression by 0% (A), 75% (B) and 100% (C) of the original sample height. Hoechst 328 was used to stain for nuclei of dead cells.

Fig. 10. Quantification of cell death at different levels of compression. The loading control corresponds to transfer to the compression chamber only, without contact with the chuck; this is closest to the actual culture conditions. 0% compression indicates contact and zero-force equilibration with the chuck of the uniaxial testing machine (TextureAnalyzer XTPlus), but no actual sample compression. 75% indicates a decrease in sample height by 75%, whereas 100% indicates theoretical total compression. The fraction of dead cells is the fraction of predominantly blue cells after Hoechst 328 staining, as compared to the total number of cells (predominantly blue + predominantly red + predominantly green). The compression experiments were repeated twice, once with starting material from a 6-well-plate culture, once with starting material from a 24-well- plate culture. Statistical significance was evaluated pairwise by comparison to the 0% compression condition. For this, linear regression with the experiment and condition (0% compression vs. loading control, 75% compression, 100% compression) as explanatory variables and the dead cell fraction as the outcome was used, and the P-value associated with the condition evaluated for significance.

Fig. 11. Quantification of the composition of the remaining viable cell fraction. The DsRed+ fraction is the fraction of red fluorescent cells compared to green + red fluorescent cells; conditions are as for Fig. 10, and evaluation based on the same images. Statistical significance was evaluated pairwise by comparison to the 0% compression condition. For this, linear regression with the experiment and condition (0% compression vs. loading control, 75% compression, 100% compression) as explanatory variables and the DsRed+ fraction as the outcome was used, and the P- value associated with the condition evaluated for significance.

Fig. 12. Uniaxial compression: Stress-Strain diagram. Four measurements were done: to 75% compression, and then to 100% compression on a total of two samples (one reconstituted from each culture condition).

Fig. 13. Young modulus as a function of polymer concentration. Eq. 4 is used for the estimation of the Young modulus from the data shown in Fig. 12; eq. 3 is used to estimate the polymer concentration from the strain shown in Fig. 12. The triangles labelled “0% compression” and “75% compression” outline the estimated polymer concentrations at which the viability quantifications for the 0% compression and 75% compression conditions were done; the grey triangle labeled “Implantation” indicates the estimated concentration of the biomaterial for the in-vivo implantation experiments (26mg/mL, see Example 1).

Fig. 14. Transplantable bone marrow niche. (A) For in vitro culture, stromal cells (OP9) are combined with hematopoietic stem and progenitor cells (HSPC, selected from bone marrow as lineage-, ckit+, Sca-1+ cells); the resulting cell mix is loaded onto collagen- coated carboxymethylcellulose microparticles (CCMs). B) During in-vitro culture, the stromal cells adhere to the scaffold and at the same time provide support to the proliferating and differentiating HSPC. C) For in-vivo implantation, the cell-loaded CCMs are slowly dehydrated to form a paste-like implantable living biomaterial. Both dehydration speed and final dehydration level are carefully controlled. D) The resulting tissue-mimicking biomaterial is injected subcutaneously for in vivo follow-up. E) Structure of a CCM. F) CCM (stained by cell impermeant Hoechst dye) along with green fluorescent stroma and red fluorescent hematopoietic compartment. G) Assignment of the different areas as scaffold, HSPCs and lineage-committed progenitors, and stromal cells (OP-9). Confocal images are linearly contrast adjusted.

Fig. 15. In-vitro co-culture of OP9 MSCs and HSPCs on CCMs. (A) Serial confocal imaging qualitatively demonstrates an increase in hematopoietic populations over the span of nearly two weeks in co-culture, for both 1:10 and 1:100 seeding densities (HSPC:MSC); green = GFP+ OP9 MSCs, red = DsRed+ HSPCs; scale bar = 100 um. Qualitative observations from imaging demonstrate large-scale structural outline of the seeded CCMs. (B) scale bar = 200 um. DsRed+ HSPCs requiring MSCs to attach and proliferate (scale bar = 100 um), very few DsRed+ cells are found at 24h with OP9 support. (C-E) focal imaging of co-seeded scaffolds at higher magnification showing HSPCs nestling within the MSC feeder layers (D, scale bar = 200 um), and 3D colonies (E), appearing throughout the scaffolds over as short as four days in culture (scale bar = 50-500 um, indicated on image). Note that in the absence of OP9 stroma (C) HSCPs are not retrieved within the scaffold at day 4 (F-G) Quantification of proliferation ratios for HSPC to OP9 in co-seeded scaffolds from confocal image z-stacks (F for 1:10 seeding ratio, G for 1:100 seeding ratio, 25 total images per “n”; n = 8 per condition, total of 200 slices). Confocal images are linearly contrast adjusted.

Fig. 16. 3D culture outcome compared to 2D controls via flow cytometry and colony forming assays. (A-C) Example flow cytometry analysis gating (Coculture in CCM scaffolds, initial seeding density KLS:OP9=1:10, analysis at 12 days. After initial gating to live cells (low DAP1 labeling due to intact membrane, isolation from counting beads, not shown), we performed a first gating using the intrinsic DsRed (HSPC and progeny) and GFP (OP9) expression (A). The DsRed+GFP- was further analyzed for Lineage and CD45 markers, allowing to isolate the Lin-CD45+ population (B). The stem and progenitor fraction was finally obtained as cKit+ cells within the Lin-CD45+ population (C). (D) Total CD45+ expansion through flow cytometry, identifying 2D and 3D cell proliferation for both the 1:10 and 1:100 seeding densities. (E) Total CD45+, cKit+ cell expansion for the same conditions, demonstrating closer similarities between the four conditions. (F) Total colony count after 7 days in methylcellulose medium, after harvesting total cells from the CCMs (after 12 days of in vitro culture).

Fig. 17. Implantation of CCM-based co-cultures. (A) After seeding, the co-cultures can be cultured in-vitro as classical microcarrier suspension cultures. (B) To prepare an implant, the material is partially dehydrated to by equilibrating to a predefined hydrostatic pressure level (DR), typically on the order of 0.2kPa (ca. 2cm water column). This is done in a specifically designed transfer tip. (C) Once equilibrated and filled with implantable co-culture biomaterial, the transfer tip is attached to a syringe and an implantation catheter (to avoid accidental intravascular injection). (D) The co-culture biomaterial is injected subcutaneously. (E) In vitro assessment of injection viability as quantified through GFP+ OP9 MSC and HSPC cell confluence before, immediately after, and 24 hours after injection (biomaterial seeded 1:100 HSPC - OP9, used at day 1 in vitro). (F) Macroscopic external view of the implant in the subcutaneous dermal tissue 12-weeks post-implantation. (G) Visibly vascularized scaffold after sacrifice, seen from the inside of the skin flap.

Fig. 18. Histology and cellular composition of implanted scaffolds. Unseeded CCMs (A-C), as well as CCMs cultured with OP9 (D-F) or with 1:10 “high” co- cultures of OP9 and KLS cells (G-I) were implanted into the dorsal skin of NSG mice, and retrieved after sacrifice at 12 weeks. Samples were processed for hematoxylin/eosin (H&E) staining (A, D, G), as well as immunohistochemistry with primary antibodies directed against GFP (B, E, H, marker for OP9 cells) and DsRed (C, F, I, marker of HSPC and progeny).

Fig. 19. Immunohistochemistry of scaffolds transplanted in vivo. Unseeded scaffolds (A- C) and HSPC/OP9-seeded scaffolds (D-I) were recovered 12 weeks after subcutaneous transplant. Paraffin sections of scaffolds were stained with anti-vWF (A, D, G, arrows indicate megakaryocytes), anti-Perilipin (B, E, H), and anti- CD31 (C, F, I), and antibodies. Scale bars are lOOpm. CD31+ vessels were quantified (J), error bars indicate mean HSD (p<0.003).

Fig. 20. Assessment of cell presence and viability during injection of compacted Cytodex3 microcarrier biomaterials; OP9 cells (GFP+, 5*106/mL) were seeded onto Cytodex3 microcarriers (40mg in 2mL) followed by partial dehydration to a interstitial pressure of 150Pa. Then, the compacted material was ejected through a 20G needle. Cell death was measured by propidium iodide staining, confluency by the GFP+ signal. Even though a significant amount of cells is lost during the injection of the Cytodex3 microcarrier, about 50% of the cells remain, the amount of remaining cells being significantly different from 0 (P=0.011).

Fig. 21. In-vivo cell transfer efficiency; Various microcarriers were seeded with OP9 cells (GFP+). For the CCM (porous, synthesis described in Example 2), 3*106 cells/mg were seeded, for the others 30*106 cells/mg to compensate for anticipated losses. Relative cell transfer efficiency was quantified from histological cuts as the number of nuclei associated with GFP+ cells.

Fig. 22 Dehydration rate measured as volumetric flow rate in CCM as function of target pressure and initial polymer concentration. A) Applied pressure is varied by varying the fluid level in the waste reservoir, initial carrier concentration 7.5mg/mL dry polymer/mL. B) Initial polymer concentration is varied, 5cm H20 of water column of aspiration pressure applied. Figure 22 shows a linear regression Y = -3.775*X + 48.59, R² = 0.74

Fig. 23. Estimated linear fluid velocity at the neck of the transfer tip (maximum expected linear average fluid velocity, 4mm neck diameter). From the data of Fig. 22.

Figure 24: A) Non-layered, compacted microcarrier culture of a non-adhesive cell type (BC1, human neural stem cells). B) Layered compacted microcarrier culture of first adhesive cell type (human foreskin fibroblast) and secondary adhesive cell type (BC1). C) Microcarrier-tissue composite (LUHMES organoids). (1) shows the diffusive membrane while (2) shows the microcarrier-tissue composite.

Figure 25: A-C) In situ re-loading of subcutaneously injected microcarrier system. A) Intact skin after re-loading of the microcarrier system with cell-emulating low diffusion carbon microparticles. B) Localized re-injection site (black) in the intact microcarrier system (encircled in white), where the dermis and epidermis have been removed. C) A magnification of the explanted microcarrier system showing the microscale distribution of the re-injected microparticles (black). D-E) Intra-bone injection of the microcarrier system. D) Fractured tibial plate at the site of injection (1) and the delivered microcarrier system (2) into the marrow space (3). E Magnification of the microcarrier system (2) integrated within the bone marrow hematopoietic component (3).

Bibliography

Beduer, A., Braschler, T., Peric, 0., Fantner, G.E., Mosser, S., Fraering, P.C., Bencherif, S., Mooney, D.J., Renaud, P., 2015a. A compressible scaffold for minimally invasive delivery of large intact neuronal networks. Adv. Healthc. Mater. 4, 301-312. https://doi.org/10.1002/adhm.201400250

Beduer, A., Braschler, T., Peric, 0., Fantner, G.E., Mosser, S., Fraering, P.C., Bencherif, S., Mooney, D.J., Renaud, P., 2015b. A compressible scaffold for minimally invasive delivery of large intact neuronal networks. Adv. Healthc. Mater. 4, 301-312. https://doi.org/10.1002/adhm.201400250

Beduer, A., Piacentini, N., Aeberli, L., Da Silva, A., Verheyen, C.A., Bonini, F., Rochat, A., Filippova, A., Serex, L., Renaud, P., Braschler, T., 2018. Additive manufacturing of hierarchical injectable scaffolds for tissue engineering. Acta Biomater. 76, 71-79. https://doi.Org/10.1016/j.actbio.2018.05.056 Bello, A.B., Park, H., Lee, S.-H., 2018. Current approaches in biomaterial-based hematopoietic stem cell niches. Acta Biomater. 72, 1-15. https://doi.Org/10.1016/j.actbio.2018.03.028 Bencherif, S.A., Sands, R.W., Bhatta, D., Arany, P., Verbeke, C.S., Edwards, D.A., Mooney, D.J., 2012a. Injectable preformed scaffolds with shape-memory properties. Proc. Natl. Acad. Sci. https://doi.org/10.1073/pnas.1211516109 Bencherif, S.A., Sands, R.W., Bhatta, D., Arany, P., Verbeke, C.S., Edwards, D.A., Mooney, D.J., 2012b. Injectable preformed scaffolds with shape-memory properties. Proc. Natl. Acad. Sci. U. S. A. 109, 19590-19595. https://doi.org/10.1073/pnas.1211516109 Blache, U., Metzger, S., Vallmajo-Martin, Q., Martin, L, Djonov, V., Ehrbar, M., 2016. Dual Role of Mesenchymal Stem Cells Allows for Microvascularized Bone Tissue-Like Environments in PEG Hydrogels. Adv. Healthc. Mater. 5, 489-498. https://doi.org/10.1002/adhm.201500795

Bourgine, P.E., Klein, T., Paczulla, A.M., Shimizu, T., Kunz, L., Kokkaliaris, K.D., Coutu, D.L., Lengerke, C., Skoda, R., Schroeder, T., Martin, L, 2018. In vitro biomimetic engineering of a human hematopoietic niche with functional properties. Proc. Natl. Acad. Sci. U. S. A. 115, E5688-E5695. https://doi.org/10.1073/pnas.1805440115 Choi, J.S., Harley, B.A.C., 2017. Marrow-inspired matrix cues rapidly affect early fate decisions of hematopoietic stem and progenitor cells. Sci. Adv. 3. https://doi.org/10.1126/sciadv.1600455 Costa, Soure, A.M. de, Cabral, Ferreira, F.C., Silva, C.L. da, 2018.

Hematopoietic Niche - Exploring Biomimetic Cues to Improve the Functionality of Hematopoietic Stem/Progenitor Cells. Biotechnol. J. 13, 1700088. https://doi.org/10.1002/biot.201700088

Croughan, M.S., Giroux, D., Fang, D., Lee, B., 2016. Chapter 5 - Novel Single-Use Bioreactors for Scale-Up of Anchorage-Dependent Cell Manufacturing for Cell Therapies, in: Cabral, J.M.S., Lobato de Silva, C., Chase, L.G., Margarida Diogo, M. (Eds.), Stem Cell Manufacturing. Elsevier, Boston, pp. 105-139. https://doi.org/10.1016/B978-0- 444-63265-4.00005-4 de la Puente, P., Muz, B., Gilson, R.C., Azab, F., Luderer, M., King, J., Achilefu, S., Vij, R., Azab, A.K., 2015. 3D tissue-engineered bone marrow as a novel model to study pathophysiology and drug resistance in multiple myeloma. Biomaterials 73, 70- 84. https://doi.Org/10.1016/j.biomaterials.2015.09.017 Fairfield, H., Falank, C., Farrell, M., Vary, C., Boucher, J.M., Driscoll, H., Liaw, L., Rosen, C.J., Reagan, M.R., 2019. Development of a 3D bone marrow adipose tissue model. Bone 118, 77-88. https://doi.Org/10.1016/j.bone.2018.01.023 Friedenstein, A.J., Latzinik, N.W., Grosheva, A.G., Gorskaya, U.F., 1982. Marrow microenvironment transfer by heterotopic transplantation of freshly isolated and cultured cells in porous sponges. Exp. Hematol. 10, 217-227.

Garg, T., Singh, 0., Arora, S., Murthy, R., 2012. Scaffold: a novel carrier for cell and drug delivery. Crit. Rev. Ther. Drug Carrier Syst. 29, 1-63. https://doi.org/10.1615/critrevtherdrugcarriersyst.v29.il.10 Henriksson, I., Gatenholm, P., Hagg, D.A., 2017. Increased lipid accumulation and adipogenic gene expression of adipocytes in 3D bioprinted nanocellulose scaffolds. Biofabrication 9, 015022. https://doi.org/10.1088/1758-5090/aa5clc Holzapfel, B.M., Hutmacher, D.W., Nowlan, B., Barbier, V., Thibaudeau, L., Theodoropoulos, C., Hooper, J.D., Loessner, D., Clements, J.A., Russell, P.J., Pettit, A.R., Winkler, I.G., Levesque, J.-P., 2015. Tissue engineered humanized bone supports human hematopoiesis in vivo. Biomaterials 61, 103-114. https://doi.Org/10.1016/j.biomaterials.2015.04.057 Hughes, C.S., Postovit, L.M., Lajoie, G.A., 2010. Matrigel: a complex protein mixture required for optimal growth of cell culture. Proteomics 10, 1886-1890. https://doi.org/10.1002/pmic.200900758

Lecarpentier, Y., Schussler, 0., Sakic, A., Rincon-Garriz, J.M., Soulie, P., Bochaton-Piallat, M.- L., Kindler, V., 2018. Human Bone Marrow Contains Mesenchymal Stromal Stem Cells That Differentiate In Vitro into Contractile Myofibroblasts Controlling T Lymphocyte Proliferation. Stem Cells Int. 2018, 6134787. https://doi.org/10.1155/2018/6134787

Leisten, I., Kramann, R., Ventura Ferreira, M.S., Bovi, M., Neuss, S., Ziegler, P., Wagner, W., Knuchel, R., Schneider, R.K., 2012. 3D co-culture of hematopoietic stem and progenitor cells and mesenchymal stem cells in collagen scaffolds as a model of the hematopoietic niche. Biomaterials 33, 1736-1747. https://doi.Org/10.1016/j.biomaterials.2011.ll.034 Maniatis, A., Tavassoli, M., Crosby, W.H., 1971. Origin of osteogenic precursor cells in extramedullary marrow implants. Blood 38, 569-575.

Mattiucci, D., Naveiras, 0., Poloni, A., 2018. Bone Marrow ‘Yellow” and “Red” Adipocytes”: Good or Bad Cells? Curr. Mol. Biol. Rep. 4, 117-122. https://doi.org/10.1007/s40610-018-0098-6 McKinney-Freeman, S.L., Naveiras, 0., Yates, F., Loewer, S., Philitas, M., Curran, M., Park, P.J., Daley, G.Q., 2009. Surface antigen phenotypes of hematopoietic stem cells from embryos and murine embryonic stem cells. Blood 114, 268-278. https://doi.org/10.1182/blood-2008-12-193888 Mcniece, I.K., Bertoncello, 1., Kriegler, A.B., Quesenberry, P.J., 1990. Colony-forming cells with high proliferative potential (HPP-CFC). Int. J. Cell Cloning 8, 146-160. https://doi.org/10.1002/stem.5530080302

Moosbrugger, C., 2002. Atlas of Stress-Strain Curves. ASM International: The Materials Information Society.

Nakano, T., Kodama, H., Honjo, T., 1994. Generation of lymphohematopoietic cells from embryonic stem cells in culture. Science 265, 1098-1101. https://doi.org/10.1126/science.8066449

Naveiras, 0., 2009. Novel determinants of the hematopoietic microenvironment in development and homeostasis - ProQuest. Harvard University, ProQuest Dissertations Publishing.

Newland, B., Welzel, P.B., Newland, H., Renneberg, C., Kolar, P., Tsurkan, M., Rosser, A., Freudenberg, U., Werner, C., 2015. Tackling cell transplantation anoikis: an injectable, shape memory cryogel microcarrier platform material for stem cell and neuronal cell growth. Small Weinh. Bergstr. Ger. 11, 5047-5053. https://doi.org/10.1002/smll.201500898

Nih, L.R., Moshayedi, P., Llorente, I.L., Berg, A.R., Cinkornpumin, J., Lowiy, W.E., Segura, T., Carmichael, S.T., 2017. Engineered HA hydrogel for stem cell transplantation in the brain: Biocompatibility data using a design of experiment approach. Data Brief 10, 202-209. https://doi.Org/10.1016/j.dib.2016.ll.069 Reinisch, A., Hernandez, D.C., Schallmoser, K., Majeti, R., 2017. Generation and use of a humanized bone-marrow-ossicle niche for hematopoietic xenotransplantation into mice. Nat. Protoc. 12, 2169-2188. https://doi.org/10.1038/nprot.2017.088 Roberts, A.S., Shetty, A.S., Mellnick, V.M., Pickhardt, P.J., Bhalla, S., Menias, C.O., 2016. Extramedullary haematopoiesis: radiological imaging features. Clin. Radiol. 71, 807-814. https://doi.Org/10.1016/j.crad.2016.05.014 Rodling, L., Schwedhelm, I., Kraus, S., Bieback, K., Hansmann, J., Lee-Thedieck, C., 2017. 3D models of the hematopoietic stem cell niche under steady-state and active conditions. Sci. Rep. 7, 4625. https://doi.org/10.1038/s41598-017-04808-0 Salvade, A., Belotti, D., Donzelli, E., D’Amico, G., Gaipa, G., Renoldi, G., Carini, F., Baldoni, M., Pogliani, E.M., Tredici, G., Biondi, A., Biagi, E., 2007. GMP-grade preparation of biomimetic scaffolds with osteo-differentiated autologous mesenchymal stromal cells for the treatment of alveolar bone resorption in periodontal disease. Cytotherapy 9, 427-438. https://doi.org/10.1080/14653240701341995 Sanchez-Aguilera, A., Mendez-Ferrer, S., 2017. The hematopoietic stem-cell niche in health and leukemia. Cell. Mol. Life Sci. CMLS 74, 579-590. https://doi.org/10.1007/s00018-016-2306-y Serex, L., Braschler, T., Filippova, A., Rochat, A., Beduer, A., Bertsch, A., Renaud, P., 2018. Pore Size Manipulation in 3D Printed Cryogels Enables Selective Cell Seeding. Adv. Mater. Technol. 3, 1700340. https://doi.org/10.1002/admt.201700340 Shafiee, A., Baldwin, J.G., Patel, J., Holzapfel, B.M., Fisk, N.M., Khosrotehrani, K., Hutmacher, D.W., 2017. Fetal Bone Marrow-Derived Mesenchymal Stem/Stromal Cells Enhance Humanization and Bone Formation of BMP7 Loaded Scaffolds. Biotechnol. J. 12. https://doi.org/10.1002/biot.201700414 Shah, N.J., Mao, A.S., Shih, T.-Y., Kerr, M.D., Sharda, A., Raimondo, T.M., Weaver, J.C., Vrbanac, V.D., Deruaz, M., Tager, A.M., Mooney, D.J., Scadden, D.T., 2019. An injectable bone marrow-like scaffold enhances T cell immunity after hematopoietic stem cell transplantation. Nat. Biotechnol. 37, 293-302. https://doi.org/10.1038/s41587-019-0017-2 Shah, S.B., Singh, A., 2017. Creating artificial lymphoid tissues to study immunity and hematological malignancies. Curr. Opin. Hematol. 24, 377-383. https://doi.org/10.1097/MOH.0000000000000356 Shih, Y.-R., Kang, H., Rao, V., Chiu, Y.-J., Kwon, S.K., Varghese, S., 2017. In vivo engineering of bone tissues with hematopoietic functions and mixed chimerism. Proc. Natl. Acad. Sci. U. S. A. 114, 5419-5424. https://doi.org/10.1073/pnas.1702576114 Song, Y., Rongvaux, A., Taylor, A., Jiang, T., Tebaldi, T., Balasubramanian, K., Bagale, A., Terzi, Y.K., Gbyli, R., Wang, X., Fu, X., Gao, Y., Zhao, J., Podoltsev, N., Xu, M., Neparidze, N., Wong, E., Torres, R., Bruscia, E.M., Kluger, Y., Manz, M.G., Flavell, R.A., Halene, S., 2019. A highly efficient and faithful MDS patient-derived xenotransplantation model for pre-clinical studies. Nat. Commun. 10, 366. https://doi.org/10.1038/s41467-018-08166-x Tanzeglock, T., Soos, M., Stephanopoulos, G., Morbidelli, M., 2009. Induction of mammalian cell death by simple shear and extensional flows. Biotechnol. Bioeng. 104, 360- 370. https://doi.org/10.1002/bit.22405

Tavassoli, M., Crosby, W.H., 1968. Transplantation of marrow to extramedullary sites.

Science 161, 54-56. https://doi.org/10.1126/science.161.3836.54 Torisawa, Y., Spina, C.S., Mammoto, T., Mammoto, A., Weaver, J.C., Tat, T., Collins, J.J., Ingber, D.E., 2014. Bone marrow-on-a-chip replicates hematopoietic niche physiology in vitro. Nat. Methods 11, 663-669. https://doi.org/10.1038/nmeth.2938

Ventura Ferreira, M.S., Bergmann, C., Bodensiek, 1., Peukert, K., Abert, J., Kramann, R., Kachel, P., Rath, B., Rutten, S., Knuchel, R., Ebert, B.L., Fischer, H., Brummendorf, T.H., Schneider, R.K., 2016. An engineered multicomponent bone marrow niche for the recapitulation of hematopoiesis at ectopic transplantation sites. J. Hematol. Oncol.J Hematol Oncol 9, 4. https://doi.org/10.1186/sl3045-016-0234-9 Weiss, L., 2008. Haemopoiesis in Mammalian Bone Marrow, in: Ciba Foundation Symposium 84 - Microenvironments in Haemopoietic and Lymphoid

Differentiation. Wiley-Blackwell, pp. 5 - 21. https://doi.org/10.1002/9780470720660.ch2 Weissman, I.L., Shizuru, J.A., 2008. The origins of the identification and isolation of hematopoietic stem cells, and their capability to induce donor-specific transplantation tolerance and treat autoimmune diseases. Blood 112, 3543-3553. https://doi.org/10.1182/blood-2008-08-078220 Xia, P., Zhang, K., Gong, Y., Li, G., Yan, S., Yin, J., 2017. Injectable Stem Cell Laden Open Porous Microgels That Favor Adipogenesis: In Vitro and in Vivo Evaluation. Acs Appl. Mater. Interfaces 9, 34751-34761. https://doi.org/10.1021/acsami.7bl3065

Yoshii, T., Sotome, S., Torigoe, L, Tsuchiya, A., Maehara, H., lchinose, S., Shinomiya, K., 2009. Fresh bone marrow introduction into porous scaffolds using a simple low- pressure loading method for effective osteogenesis in a rabbit model. J. Orthop. Res. Off. Publ. Orthop. Res. Soc. 27, 1-7. https://doi.org/10.1002/jor.20630

Claims

Claims

1. A process for direct in-vitro to in-vivo transfer of a living microcarrier culture by controlled dehydration into a porous, viscoelastic material capable of cell-protection during delivery, characterized in that

- the final pressure of the interstitial fluid in the compacted implant for injection is between 20 Pa and 2kPa, preferentially 50 Pa to lkPa, and even more preferentially between 100 Pa and 800 Pa below atmospheric pressure

- the flow rate in the part of material during dehydration remains between O.Olmm/s and lOcm/s, more preferentially between O.lmm/s and 5cm/s, and even more preferentially between 0.5mm/s and lOmm/s.

2. A device for performing the process of controlled dehydration of a living microcarrier culture of claim 1, wherein the device is arranged for guaranteeing a set final precision in the range of 20Pa to 2kPa to within a precision of 500 Pa, preferentially 200 Pa, even more preferentially 100 Pa and most preferentially 20Pa, irrespective of the initial cell culture volume, and a local flow rate between 0 and lOcm/s in the material to be injected to within a precision of +/- 50%, preferentially +/- 20%, and even more preferentially +/- 10%, under aseptic conditions.

3. A device consisting of the dehydration device and a transfer device that on the one hand allows connection to the dehydration device, and on the other plug- or screwing connection to an injection device such as a syringe and needle.

4. A composition comprising a viscoelastic fluid comprising essentially optimized cell carriers for the process described in claim 1, in that: a) The cell carriers are partially elastically and reversibly compressed, preferentially in the range between 2.5% and 100% of the fully swelled volume, more preferentially between 5% and 80% and most preferentially between 10% and 60% of the fully swelled volume at the interstitial fluid pressure for in-vivo delivery, stabilizing said pressure against small variations of hydration b) The cell carriers are cell protective in that partial compression to a range of 2.5% and 100% of the fully swelled volume, more preferentially between 5% and 80% and most preferentially between 10% and 60% of the fully swelled volume at the interstitial fluid pressure for in-vivo delivery does not cause more than an addition 50%, preferentially 20% and most preferentially 10% of cell mortality. c) The average pore size in the material for implantation at the interstitial pressure chosen for delivery is between 20 micrometers and 1cm, preferentially between 40 micrometers and 1mm, and even more preferentially between 50 micrometers and 500 micrometers. d) The cell carriers are sufficiently hydrophilic to retain aqueous liquid rather than letting the ambient atmosphere penetrate during dehydration, resulting at least 50%, preferentially 80% and even more preferentially 90% of the interstitial space between and possibly within microcarriers if they are porous being liquid rather than gas phase.

5. A composition according to claim 4, wherein the viscoelastic fluid is mechanically sufficiently strong to sustain a given shape in-vivo, by presenting less than 30%, preferentially less than 20% and even more preferentially less than 10% change in shape as measured by length:width aspect ratio within 3 weeks, preferentially 3 months, and even more preferentially 6 months.

6. A microcarrier compatible with layered adhesive co-culture in that it provides specific adhesion to a first cell type, but not a second cell type, where in addition the second cell type will adhere on the first cell type.

7. A microcarrier according to claim 6, wherein at least one of the two cell types is a human cell.

8. A microcarrier according to claim 6 or 7, wherein at least part of the final amount of the first or second cell types is provided after a first implantation, by re- injection possibly by keeping the integrity of the living microcarrier system.

9.. A microcarrier according to any one of claims 6 to 8, wherein at least part of the second cell type, which directly associates with the delivered adherent cell type, is provided by the host upon injection.

10. A device for performing the process of controlled dehydration of a living microcarrier culture described in claim 2, where after initial dehydration, medium is slowly replenished, maintaining continuous flow at the set pressure for several seconds, preferentially several minutes and even more preferentially several days, the culture being conditioned by said flow

11. A device according to claim 2, wherein additionally the compacted living microcarrier culture is separated from the flow by a diffusive membrane permitting exchange of nutrients and possibly proteins, but limiting the flow in the compacted microcarrier material to below 10 micrometers/s, preferentially below 1 micrometer/s, even more preferentially to below lOOnm/s and most preferentially below lOnm/s, whereas the flow rate on the supply side of the diffusive membrane is at least 2x, preferentially lOx and most preferentially lOOx superior to the one in the scaffold material.

12. A device according to claim 1, where a living microcarrier culture is obtained by adding or layering an organoid, tissue, or cell mass on top of compacted microcarriers, and where during dehydration or the additional time of incubation as in claim 10, the organoid, tissue or cell mass is drawn at least partially into the compacted microcarrier mass by the dehydration pressure in the range of 20Pa to 2kPa, preferentially lOOPa to 1.5kPa, and most preferentially 300Pa to lkPa, to form a microcarrier-tissue composite during dehydration or extended culture as in claim 10.

13. The microcarrier-tissue composite obtained by the process described in claim 1

14. A device for performing the process of controlled dehydration of a living microcarrier culture described in claim 10, where a by-product is recovered from the flow.

15. A device for performing the process of controlled dehydration of a living microcarrier culture described in claim 2, where the flow-limiting resistance is a self-regulating porous material that upon attempted increase in flow decreases its pore size and thus increases the flow resistance, and visa-versa expands upon a decrease of flow-rate to counteract the decrease

16. A device for performing the process of controlled dehydration of a living microcarrier culture described in claim 15, where the microcarriers are designed in such a way as to self-assemble the porous flow-regulating material, by possessing a Young modulus above such that at the desired limiting flow rate, the compression becomes sufficiently important to limit flow rate increase to no more than 100% regardless of what pressure up to the final desired interstitial pressure is applied

17. A device for performing the process of controlled dehydration of a living microcarrier culture described in claim 2 that can sterilized by one of the usual means such as gamma- sterilization, ethylene oxide sterilization or autoclave sterilization.

18. A transfer device as part of the overall device described in claim 3, designed such that it maintains the compacted microcarriers when lifting the transfer device off the capillary conductor (or filter membrane) of the dehydration device, achieving this by being conical with cone opening angles between 0° and 180°, preferentially 1° to 90°, more preferentially between 2° and 45°, even more preferentially between 3° and 30°, and most preferentially between 5° and 20°.

19. A dehydration and transfer device as described in claim 3 with a filter membrane interposed between transfer device and dehydration device

20. A transfer device where the tip opening performs regulation of the fluid flow rate by having a tip opening diameter is in between about 20% of the fully expanded microcarrier size to a maximum of about 5cm, more preferentially between 50% of the average fully expanded microcarrier size and 1cm, and even more preferentially between 100% of the average fully expanded microcarrier size and 0.5cm.

21. A living microcarrier suspension as prepared by claim 1, where one or several additional functional elements are added to the suspension such as a visosing agent, increasing the viscosity of the interstitial fluid in the range of 2x to 10⁶x, more preferentially lOx to 10⁴x, and/or pharmacologically active ingredients such as growth factors and more particularly VEGF, PDGF and/or FGFs, cells secreting such growth factors either spontaneously or by genetic modification, cytokines or other hematopoiesis-promiting, immune-modulating or other factors inducing cell growth and/or differentiation and more particularly erythropoietin, thrombopoietin, Flt3-ligand, SCF, dimethyloxaloylglycine, Stem regenin 1, notch ligands such as D114, NGF and/or BMPs.