WO2022096884A1

WO2022096884A1 - Method of culturing cells

Info

Publication number: WO2022096884A1
Application number: PCT/GB2021/052859
Authority: WO
Inventors: Peter Richard COOK
Original assignee: Oxford University Innovation Limited
Priority date: 2020-11-06
Filing date: 2021-11-04
Publication date: 2022-05-12
Also published as: GB202017551D0

Abstract

Methods of culturing cells are disclosed. In one arrangement, a cell holding structure is used that has a plurality of openings. The cell holding structure contains an aqueous cell growth environment. The cell growth environment is submerged in a cover liquid that is immiscible with water and retains water within the cell growth environment by interfacial tension at interfaces between the water and the cover liquid across at least a subset of the openings. Living cells are provided in the cell growth environment. The cells are cultured by providing a flow of cell culture media into and out of the cell growth environment through at least one of the openings while the cell growth environment is submerged and confined by the cover liquid.

Description

METHOD OF CULTURING CELLS

The present disclosure relates to culturing cells and is particularly applicable to growing tissue for medical use and for manufacturing food like artificial meat.

Growing tissue outside of the body for use in replacing damaged, missing or diseased tissue is an active area of research with great medical potential. There are various practical challenges to be overcome, including how to reliably provide suitable flows of nutrients and waste to and from cells in the growing tissue, as well as how to build complex tissue structures. Similar practical considerations apply for manufacturing artificial meat. Existing methods tend to be complex and expensive, or lack flexibility.

It is an object of the present disclosure to at least partly address one or more of the challenges in the prior art discussed above and/or to improve cell culturing generally in the area of tissue growth for medical purposes and/or food production.

According to an aspect of the invention, there is provided a method of culturing cells, comprising: providing a cell holding structure having a plurality of openings, wherein the cell holding structure contains an aqueous cell growth environment, and the cell growth environment is submerged in a cover liquid that is immiscible with water and retains water within the cell growth environment by interfacial tension at interfaces between the water and the cover liquid across at least a subset of the openings; providing living cells in the cell growth environment; and culturing the cells in the cell growth environment by providing a flow of cell culture media into and out of the cell growth environment through at least one of the openings while the cell growth environment is submerged and confined by the cover liquid.

Thus, a method is provided in which cells are cultured in a cell holding structure that is submerged in a cover liquid immiscible with water. The cell holding structure has self-sealing openings that allow cell culture media to be fed through the structure and selfseal otherwise. Water is retained in the cell growth environment by interfacial tension associated with interfaces spanning the openings. This approach allows cells to be grown while being robustly protected from the outside environment. The approach is also flexible in that the size and shape of the cell holding structure can be varied easily without changing the mechanism of operation. In an embodiment, the cell holding structure comprises a plurality of detachable cell holding sub-structures, each cell holding sub-structure comprising a plurality of openings and containing a respective portion of the aqueous cell growth environment. The cell holding structure may thus be modular. In some embodiments, living cells are provided in some or all of the cell holding sub-structures before the cell holding substructures are moved into contact with each other. This approach provides enhanced flexibility, allowing different cells or mixtures of cells to be provided in different parts of the assembled cell holding structure.

In an embodiment, water is held in the cell holding sub-structures by interfacial tension at interfaces between the water and surrounding medium during moving of the cell holding sub-structures into contact with each other. Each respective portion of the final aqueous cell growth environment is thus protected during the assembly of the cell holding structure.

In an embodiment, each of at least a subset of the interfaces between the water and surrounding medium is replaced by a continuous aqueous liquid connection between the respective cell holding sub-structures when the cell holding sub-structures contact each other. Thus, interfaces at the openings retain water by interfacial forces while outer surfaces of cell holding sub-structures are exposed to a surrounding cover liquid, and then convert to providing a channel by which flows of aqueous liquid can be driven through a cell growth environment spanning multiple cell holding sub-structures once those cell holding sub-structures have been brought into contact with each other. No special procedures or fine manufacturing steps are needed to fluidically connect the aqueous phases in the different modules (cell holding sub-structures) to each other. The modules simply need to be moved into contact with each other.

In an embodiment, the cell holding sub-structure comprises openings in an upper surface and a lower surface to allow vertical stacking. In an embodiment, the cell holding sub-structure comprises openings in a lateral surface to allow horizontal stacking. The cell holding sub-structures can thus be stacked together flexibly to create a wide range of shapes of tissue and/or distribution of cell types within the tissue.

In an embodiment, the cell holding sub-structures contact each other via respective planar surfaces that each contain plural openings. Plural openings in the same planar surface allows multiple corresponding flows of cell culture media to be provided. This makes it easier to ensure that there is a flow of cell culture media sufficiently close to all of the cells to ensure that the cells can develop normally. The plural openings may also facilitate connection of different modules together by helping to ensure that there is a sufficiently large number of overlapping openings at the interface to allow a desired flow of cell culture media to pass, without requiring onerous alignment procedures between openings. The openings may even be distributed at random, as might conveniently be the case where the openings are formed by pores in a porous material.

Embodiments of the disclosure will now be further described, merely by way of example, with reference to the accompanying drawings, in which:

Figure l is a schematic side sectional view depicting flow of cell culture media through a cell holding structure submerged in a cover liquid;

Figure 2 is a schematic side sectional view depicting flow of cell culture media through a cell holding sub-structure submerged in a cover liquid;

Figure 3 is a schematic side sectional view of a cell holding sub-structure comprising a substantially planar member with a hole extending through the plane of the planar member;

Figure 4 is a schematic side sectional view of a cell holding sub-structure of the type depicted in Figure 3 with a radially inward facing wall defining the hole of the planar members comprising one or more protrusions;

Figure 5 is a top view of a cell holding sub-structure of the type depicted in Figure 3 with a circular hole;

Figure 6 is a top view of a cell holding sub-structure of the type depicted in Figure 3 with a square hole;

Figure 7 depicts units isolated from square holes in cell holding sub-structures before assembly together;

Figure 8 depicts the units of Figure 7 assembled together;

Figure 9 depicts a plurality of separated cell holding sub-structures suitable for forming a vertical stack;

Figure 10 depicts a stack assembled from the cell holding sub-structures of Figure

9; Figure 11 is a schematic side sectional view of two laterally spaced stacks showing hydrostatically driven flow of cell culture media from one stack to the other stack;

Figure 12 is an image of a stack of washers acting as cell holding sub-structures containing cell culture media in the axial holes;

Figure 13 is an image of a stack comprising three washers with spacers between a lowermost washer and the middle washer;

Figures 14-17 are images depicting driving of a flow of cell culture media between different stacks via a bridge between the two stacks;

Figures 18-20 are images depicting feeding of cell culture media between different stacks via a flow conduit provided beneath the stacks;

Figure 21 is an image of a stack comprising stainless steel washers and a tubular filter paper layer;

Figures 22-24 are images of a stack comprising stainless steel washers sandwiching four layers of filter paper;

Figures 25 and 26 are images of a stack comprising two layers of hydrogel;

Figure 27 is a schematic side and top view of a stack of two cell holding substructures filled with cell culture media and overlaid with an FC40 cover liquid.

Figure 28 is a schematic side sectional view of an experimental configuration for validating operation of an example cell holding structure;

Figure 29 is an image of a configuration of the type shown in Figure 28, viewed from above, prior to starting of a flow through the cell holding structure;

Figure 30 is an image of the configuration of Figure 29, viewed from the side, 30 minutes after the flow has started;

Figures 31-40 are images depicting stages in an illustrative experiment using the configuration of Figures 29 and 30; and

Figure 41 depicts cell growth in different modules (cell holding sub-structures). Each chamber in the mesh is 400 x 400 x 340 pm (volume ~50 nl). Phase-contrast images of typical chambers at the center of meshes are shown; no attempt is made to re-image the same focal planes through identical chambers, as doing so in these 3D structures is challenging, (i) Squares of nylon mesh (10 x 10 mm) are dipped successively in ice-cold Geltrex containing HEKs (10⁵/ml) to fill chambers, then FC40 at 37°C (10 s) to set the gel, and finally into medium in which they float freely. After culture overnight, most chambers contain a few single cells (some in and other out of focus), (ii) Control: one mesh remains unstacked and floats freely as clones grow for 14 d. (iii) Stack. Cartoon: stack structure (paper base, 12 x 10 mm). Left: photograph of stack in dish filled with FC40. The stack is fed with 75 pl medium (roughly the free volume in the stack of 5 meshes) on days 1, 3, and 5 by pipetting on to the paper base; added medium passes quickly up the stack, and an equal volume is removed from the cap. On day 7, the stack is disassembled, and central chambers in the mesh originally in the center of the stack imaged. The stack is now reassembled, and fed daily for another 7 d; finally, the stack is disassembled, and central chambers in the stack re-imaged. In general, colonies appear larger than controls, but it is difficult to compare volumes precisely due to differences in 3D environments and procedures (e.g., some colonies grow out of gels in controls, while others are tom apart during unstacking).

The present disclosure relates to methods of culturing cells.

In one class of embodiment, the method comprises providing a cell holding structure 8 as exemplified in Figure 1. The cell holding structure 8 has a plurality of openings. In the example of Figure 1, the cell holding structure 8 comprises openings that are relatively small in comparison with the size of the cell holding structure 8. The cell holding structure 8 may, for example, completely or partially comprise a porous material and pores of the porous material may provide all or a portion of the openings of the cell holding structure 8. In an embodiment, the openings have characteristic dimensions of about fifty to a few hundred microns and are spaced apart roughly every fifty to a few hundred microns on average. In some embodiments, a largest dimension of each opening 8 is on average less than 10%, optionally less than 5%, optionally less than 1%, optionally less than 0.1% of a largest dimension (e.g. a length) of the cell holding structure 8. The cell holding structure 8 in the example of Figure 1 has many such small openings. The openings are not therefore depicted in Figure 1.

The cell holding structure 8 contains an aqueous cell growth environment. The cell growth environment is submerged in a cover liquid 2. An interface between the cover liquid 2 and the outside environment 4 (e.g. air) is labelled “5”. The cover liquid 2 retains water within the cell growth environment by interfacial tension at interfaces between the water (in the aqueous cell growth environment) and the cover liquid 2 across at least a subset of the openings. The interfaces thus span the openings in the cell holding structure 8. The interfaces may be concave, convex, or flat. The cover liquid 2 prevents water from leaking out of the openings in the cell holding structure 8 in an unwanted way. The cover liquid 2 also protects the cell growth environment from contamination (by, for example, microbes or unwanted chemicals) and/or reduces evaporation from the cell growth environment.

The method further comprises providing living cells in the cell growth environment in some or all of the cell holding sub-structures. The cells may be provided before the cell holding structure 8 is submerged in the cover liquid 2 and/or while the cell holding structure 8 is submerged in the cover liquid. In the latter case, media containing cells might be dispensed from the tip of a dispensing tube or pipet submerged in the cover liquid 2, through an aqueous bridge between the tip and media in an opening or pore, to a cell holding structure 8.

The method further comprises culturing the cells in the cell growth environment by providing a flow 12 of cell culture media into and out of the cell growth environment through at least one of the openings. The flow into or out of the cell growth environment may be directed into the cell growth environment along a conduit 14 (which may be a microfluidic conduit), along a tube, or along porous material (e.g. a filter paper track or similar formed along a bottom surface of a container 6 containing the cell holding structure 8 and cover liquid 2). In the example shown, the flow into the cell growth environment is provided along a flow conduit 14 and the flow out of the cell growth environment is driven by buoyancy forces that cause globules of water to form, grow and eventually leave the cell growth environment via openings in an upper surface of the cell holding structure 8 (schematically shown by arrows). The flow 12 is provided while the cell growth environment is submerged and confined by the cover liquid 2. The flow 12 may be driven by a pressure gradient along the flow conduit 14. The pressure gradient may be applied predominantly using hydrostatic pressure or by a powered pumping device or by differences in Laplace pressure. An example implementation of driving the flow using hydrostatic pressure is depicted in Figure 11 and described in further detail below.

The cell culture media (which may also be referred to as cell culture medium, growth media/medium or culture media/medium) comprises a liquid containing components (e.g. various proteins/food) for supporting survival and/or proliferation of cells. Examples include Dulbecco's Modified Eagle Medium (DMEM) or Roswell Park Memorial Institute medium (RPMI) with added serum, for example around 10% serum, for example around 10% Fetal Bovine Serum (FBS). Many nowadays use serum-free media, where the FBS or other animal derived substances are not used. Cell culture media can in general be a liquid or a gel (which typically contains liquid). For example, agar-plates for bacteria comprise a gel formed from a liquid medium. Mostly, however, eukaryotic cells are grown in liquid cell culture media. The liquid cell culture media is a source of energy (often in the form of glucose), amino acids, vitamins and some trace elements. Supplements may be added to those media, such as FBS, which supports a breadth of different cells. The precise composition of FBS is not known. FBS is not a chemically defined mix because it comes from a biological source. FBS is known, however, to contain at least hormones, lipids, proteins, and growth-factors which all regulate cell behaviour - i.e., they enable cells to grow and duplicate, regulate membrane physiology, etc. In many instances it would be beneficial to avoid using FBS. This is because 1) FBS is undefined and therefore variable, and 2) FBS comes from animals and could carry pathogens. The latter factor is important for industries that produce tissues and organs for transplantation, where it may be necessary to ensure that no components used in a whole manufacturing pipeline have animal origin. As a result, alternative supplements are now available which are either fully chemically-defined and animal-free or, if not animal-free, at least better defined than serum.

In some embodiments, the cover liquid 2 comprises a fluorocarbon such as FC40. FC40 is a transparent, bioinert, and fully fluorinated liquid of density 1.8555 g/ml that is widely used in droplet based microfluidics. It can also act as a sterility barrier around the cell holding structure 8.

In some embodiments, the cell holding structure 8 comprises a plurality of detachable cell holding sub-structures 10. The cell holding sub-structures 10 are thus not integral with each other for at least a portion of the method (e.g. when they are first brought into contact with each other). Eight such sub-structures 10 are shown in the example of Figure 1. Each cell holding sub-structure 10 comprises a plurality of openings. The openings may be configured in any of the ways described above for the openings of the cell holding structure 8. Each cell holding sub-structure 10 contains a respective portion of the aqueous cell growth environment of the cell holding structure 8. In embodiments of this type, the cell holding structure 8 may be built by moving initially separated cell holding sub-structures 10 into contact with each other. An example of such an individual cell holding sub-structure 10 is depicted in Figure 2. In certain cases, some of the cell holding sub-structures 10 may contain cell culture medium while not containing cells. In some embodiments, some of the cell holding sub-structures 10 contain cell culture medium and no living cells and some of the cell holding sub-structures 10 contain cell culture medium and living cells.

The cell holding sub-structures 10 may be referred to interchangeably as substructures or modules. The cell holding structure 8 may be referred to interchangeably as a stack in embodiments where the cell holding structure 8 is formed from a plurality of the cell holding sub-structures 10 (by stacking the sub-structures together).

In some embodiments, the cell holding structure 8 and/or each of one or more of the cell holding sub-structures 10 comprises a porous body. The porous body may, for example, comprise a matrix, scaffold, polymer or hydrogel formed from units of sugar, carbohydrate, amino-acid, or hydrocarbon residues. The porous body may thus comprise cellulose or other paper-based materials derived from plants or bacteria, starch-based materials, as well as man-made polymers. For the production of organs, the porous material may be biocompatible and biodegradable. For the production of foods like artificial meat, the porous body may be edible and biodegradable.

Living cells may be provided in the cell holding sub-structures 10 before the cell holding sub-structures 10 are moved into contact with each other. In embodiments of this type, the living cells may be cultured in the cell holding sub-structures 10 before the cell holding sub-structures 10 are moved into contact with each other. The culturing may be performed by providing a flow 12 of cell culture media (e.g. driven by a pressure gradient) into and out of each cell holding sub-structure 10 through at least one of the openings in each cell holding sub-structure 10 while the cell holding sub-structure 10 is submerged by the cover liquid 2. The flow of cell culture media into and out of each cell holding substructure 10 during this phase may occur via any of the mechanisms discussed above for flow of the cell culture media through the cell holding structure 8.

Water may be held in the cell holding sub-structures 10 by interfacial tension at interfaces between the water and surrounding medium during moving of the cell holding sub-structures 10 into contact with each other. The cell holding sub-structures 10 may be moved into contact with each other while the cell holding sub-structures 10 are submerged in the cover liquid 2. Alternatively or additionally, the cell holding sub-structures 10 may be brought into temporary contact with other media such as air (for a short period of time). The surrounding medium referred to above may thus comprise the cover liquid 2 or other media such as air or other liquids. If passing through air, the air would typically bubble off once aqueous continuity between adjacent cell holding sub-structures 10 is established. If passing through cover liquid 2, the cover liquid 2 would typically merge with the rest of cover liquid 2 once aqueous continuity between adjacent cell holding sub-structures 10 is established.

Each of at least a subset of the interfaces between the water and surrounding medium may be replaced by a continuous liquid connection between the respective cell holding sub-structures 10 when the cell holding sub-structures 10 are brought into contact each other. Each liquid connection that is made in this way provides a new route for cell culture media to be driven through the system. All of the respective portions of the aqueous cell growth environment in the different cell holding sub-structures 10 may thus be brought into continuous liquid communication with each other via at least a subset of the openings in the cell holding sub-structures 10 (i.e., those openings that connect together the respective cell holding sub-structures 10).

In some embodiments, each of one or more of the cell holding sub-structures 10 comprises openings in an upper surface and a lower surface to allow vertical stacking of the cell holding sub-structures 10 to form the cell holding structure 8. This configuration would be compatible with the example of Figure 1, the openings in the upper and lower surfaces allowing the aqueous phase in the cell holding sub-structures 10 in the lowermost layer to be connected fluidically to the aqueous phase in the cell holding sub-structures 10 in the uppermost layer.

In some embodiments, each of one or more of the cell holding sub-structures 10 comprises openings in a lateral surface to allow horizontal stacking of the cell holding sub- structures 10 to form the cell holding structure 8. This configuration would be compatible with the example of Figure 1, the openings in the lateral surfaces allowing media in all four cell holding sub-structures 10 in the uppermost layer to be connected fluidically to media in each other directly and allowing all four cell holding sub-structures 10 in the lowermost layer to be connected fluidically in this way to each other directly. Then, aqueous phases in all eight cell holding sub-structures would be connected fluidically to each other directly.

The cell holding sub-structures 10 may thus be stacked vertically and/or horizontally. To achieve close packing or complete space filling (neglecting the openings), the cell holding sub-structures 10 may have shapes that tessellate with each other when viewed horizontally and/or vertically. For example, the cell holding sub-structures 10 may have square or rectangular cross-sections when viewed horizontally and/or vertically.

In an embodiment, two or more of the cell holding sub-structures 10 contact each other via respective planar surfaces that each contain plural openings. In the example of Figure 1, it can be seen that an upwards facing planar surface of each of the cell holding sub-structures 10 in the lowermost layer contacts a respective downwards facing planar surface of a respective cell holding sub-structure 10 in the uppermost layer. Each of these planar surfaces contains plural openings. The openings may be highly numerous so as to ensure that many of the openings will overlap with each other when the respective cell holding sub-structures 10 are brought into contact with each other and thereby contribute to establishing the continuous aqueous communication between the different cell holding sub-structures 10. The openings may even be distributed at random over the planar surfaces. As mentioned above, the openings may have characteristic dimensions of several hundred microns and be spaced apart from each other on average by several hundred microns.

Providing many liquid connections between the different cell holding substructures 10 may assist with efficient cell culturing. A key requirement for any 3D culture is efficient feeding of growing cells/tissues and removing waste. As a general rule, cells in our bodies are never more than -200 microns from the vasculature, which defines the length scale over which diffusion or extra- vascular flows work in practice. If a body of cells is not fed continuously and waste removed, some cells die - often the ones in the middle. For example, induced pluripotential human stem cells can grow in static cultures into a roughly spherical ball of many cells, an ‘organoid’. Cells in such organoids can further develop into cells that express many of the markers characteristic of brain cells. However, cells in the centre of the ball often become necrotic and die when such ‘brain organoids’ become more than a few hundred microns in diameter. The openings in the different cell holding sub-structures 10 and the internal (e.g. porous) structure of the cell holding sub-structures 10 may thus be configured to ensure that the flow of the cell culture media adequately simulates performance of natural vasculature, thereby ensuring the cells are properly fed, waste is properly removed, and that cells do not die. When the cell holding sub-structures 10 are assembled together (e.g. which may be done to form a larger body of tissue or an organ), it is desirable to ensure that the cells continue to be fed properly and that waste is removed properly. Exit openings on one module therefore need to allow flows through input pores in the next module in the stack. These flow requirements may be beneficial in other ways also. First, for example, in developing embryos there is a requirement for efficient cell-to-cell communication during tissue formation. ‘Morphogens’ and other signalling molecules diffuse between cells (or are transferred between contacting cells without getting out into the external liquid) to create ‘morphogen gradients’ that drive differentiation depending on morphogen concentration - and these signals are interpreted by the cells so they differentiate in different ways to give different tissues. It is a general rule that in embryos these gradients act over short distances (i.e., over tens to hundreds of cells). This again translates to distances of about 50 microns or a few hundreds of microns, which reinforces the desirability of providing flows that are within this distance of the cells being cultured.

In some embodiments, two or more of the cell holding sub-structures 10 contain biological cells of different types relative to each other so that the culturing of cells produces a structure of cells of different type in different regions. This approach to growing tissue may be convenient because the conditions that drive differentiation of different kinds of tissue are often known individually. It may then be efficient to first grow the different cell types separately (using different media), and then assemble them in a subsequent step to provide a large unit of tissue. For example, to make a simple artificial skin, two layers may be grown separately (e.g., fibroblasts and keratinocytes, each in their own media), stacked together in a subsequent step, and then grown between a reservoir containing media suitable for fibroblasts at the bottom, and an air-filled reservoir at the top (so keratinocytes at the top can make keratin and a dry flaky ‘skin’ at the waterair interface). For example, to make a simple artificial steak, bovine myoblasts (which will differentiate into muscle) and fat cells (which will give taste and texture) may be grown separately, and then stacked on top of each other so that they can continue to grow and develop into a complex tissue. Note that it may prove beneficial to add an extra cell type to the layer containing endothelial cells (when making artificial skin), and in the layer containing myoblasts (when making artificial meat). Thus, it is known that flow promotes angiogenesis in vivo in the mouse retina, and enhances vascularisation (and so maturation) of kidney organoids in vitro. Consequently, seeding endothelial cells from arteries or veins with others in the flowing environment of cell holding sub-structure 10 may lead those endothelial cells to develop naturally into a tubular network - a primitive ‘vasculature’ - through which flow 12 could pass.

In Figures 1 and 2, the cell holding structure 8 and the cell holding sub-structures 10 are static. However, when mass-producing large quantities of skin or artificial meat, for example, it would be attractive to operate a production line that works continuously, in which a cell holding structure 8 and cell holding sub-structures 10 are mobile and dragged through the cover liquid 2. Then, one can imagine using, for example, two cell holding sub-structures 10 that were long thin strips (containing, for example, myoblasts and fat cells, respectively) that were dragged through the cover liquid 2 separately, and then stacked on top of each other to give one cell holding structure 8 that was further dragged through the cover liquid as the two cell types continued to grow and form a tissue. Optionally, the two cell types could be rolled up at the end to give, for example, the equivalent of a filet mignon. In such cases, the two layers could be interleaved with other layers that provided additional flavourings and textures.

Figure 3 depicts an alternative embodiment in which each cell holding substructure 10 comprises a substantially planar member with a hole 16 that extends through the plane of the planar member. In the example of Figure 3, the plane of the planar member is horizontal. The hole 16 in this case thus extends continuously through the cell holding sub-structure 10 in a vertical direction, opening out both into an uppermost surface 18 and lowermost surface 20 of the planar member. The planar member may be solid (non-porous) or porous. The hole 16 may be empty or a porous material (e.g. a hydrogel) may be held in the hole 16. The cell holding sub-structure 10 may thus comprise a combination of a solid, non-porous member and a porous member held by the solid, non- porous member. In some embodiments, as exemplified in Figure 4, a radially inward facing wall 24 defining the hole 16 in the planar member comprises one or more protrusions. The protrusions increase the surface area of the inward facing wall 24 and may take various forms, include ridges, ledges, threads or surface roughening. The increase in surface area provided by the protrusions may improve holding of a porous material such as a hydrogel in the hole 16.

Figures 5 and 6 are example top views of a cell holding sub-structure 10 of the type depicted in Figure 3. An outer peripheral surface 22 is circular in the examples shown, but other shapes are possible, including tessellating shapes such as squares or rectangles. The holes 16 may also have various shapes. Examples of circular and square holes 16 are shown respectively in Figures 5 and 6. It may be desirable for the holes 16 to have shapes that tessellate in embodiments where cells are grown in structures (e.g. hydrogels) held in the holes 16. These units of structure may be isolated from the outer solid portions of the cell holding sub-structures 10 and stacked together as depicted in Figures 7 and 8. Figure 7 schematically shows units 11 isolated from cell holding sub-structures 10 with square holes 16 of the type depicted in Figure 5. Figure 8 schematically shows the units 11 of Figure 7 stacked together in the horizontal plane (with fluidic connections being made at the lateral surfaces to create a fluidically continuous aqueous cell growth environment).

As exemplified in Figures 9 and 10, the cell holding sub-structures 10 may be stacked on top of each other with the holes 16 of two or more of the cell holding substructures 10 aligned with each other vertically to form at least a portion of the aqueous cell growth environment. Figure 9 depicts a plurality of separated cell holding substructures 10 suitable for forming a stack. Figure 10 depicts the cell holding sub-structures 10 stacked together. If done while the cell holding sub-structures 10 are submerged in the cover liquid 2, an aqueous cell growth environment held in each hole (e.g. in a hydrogel or other scaffold material) will form one or more interfaces between water and cover liquid interfaces on an upper side 32 of the hole 16 and on a lower side 34 of the hole 16 (as labelled in Figure 9). When the cell holding sub-structures 10 are brought into contact with each other, the interfaces between different liquids on the upper and lower sides 32, 34 are replaced by fluidically continuous connections between different cell holding substructures 10, thereby created a fluidically continuous cell growth environment spanning multiple cell holding sub-structures 10 in a region 36 formed from the holes 16 of the multiple cell holding sub-structures 10. In embodiments where the outer parts of the cell holding sub-structures 10 are also porous, the cell growth environment may extend also into the outer parts.

We have seen that if a body of cells is not fed continuously and waste removed, some cells die, for example, ones in the middle of a ‘brain organoid’ that is several hundreds of microns in diameter. Such organoids can be grown to a larger diameter without necrosis if the culture media is stirred vigorously to ensure continuous supply of nutrients and removal of waste. A module such as the one shown in Figure 3 (with a planar member that forms a solid ring around the hole 16) can be used to grow such organoids, for example in the middle layer in a 3 -module stack. The bottom layer in the stack would act as a reservoir that is connected to an input flow conduit 14; it might consist of a cell-free and porous cell holding sub-structure 10 without an internal hole 16 that has an upper surface to which cells do not adhere (e.g., a pre-wetted porous filter disc made of PTFE) and which is connected to an input flow conduit 14. The top layer in the stack would act as an upper reservoir much like that in Figure 3 with a planar member forming a solid ring around hole 16. Upward flow of media from the input flow conduit 14 to the upper reservoir would continuously supply fresh media to the developing organoid and remove of waste without stirring, so the organoid could grow to a larger diameter without internal necrosis.

Figure 11 schematically shows one way of using hydrostatic pressure to provide a pressure gradient in a conduit 14 and thereby drive the flow of cell culture media through the cell growth environment. In this example, two vertical stacks of the cell holding substructures 10 are provided. The leftmost stack is lower than the rightmost stack. The rightmost stack forms the cell holding structure 8 and contains the living cells of interest. The leftmost stack provides a reservoir of cell culture media. The leftmost stack could therefore be formed from structures other than the cell holding sub-structures 10. Cell culture media 40 is input to the leftmost stack (e.g. into central axial holes of the cell holding sub-structures 10 where the cell holding sub-structures 10 are of the type described above with reference to Figures 3-10). The height of cover liquid 2 above the leftmost stack is greater than the height of cover liquid 2 above the rightmost stack. The cover liquid 2 is denser than water in this example. A greater pressure is thus exerted on the cell culture media in the leftmost stack than on the water in the rightmost stack. This results in a pressure gradient in the conduit 14 that drives flow of the cell culture media from the left to the right, thereby driving the cell culture media through the cell holding structure 8. Excess water forms globules at the top of the rightmost stack that will grow and eventually detach from the rest of the body of liquid and rise to the surface of the cover liquid 2.

In some embodiments, the flow of cell culture media is varied to provide a pulsatile flow. A pulsatile flow may be desirable to provide a more realistic simulation of conditions in the human body. Where hydrostatic pressure is used to provide the pressure gradient driving the flow, a pulsatile flow may conveniently be provided by mechanically rocking the cell holding structure 8 and cover liquid 2 (e.g. by rocking a container containing the cell holding structure 8 and cover liquid 2). The rocking causes a pulsatile variation of the height of the cover liquid over the cell holding structure 8 and/or flow conduits leading to the cell holding structure 8 and/or reservoirs of cell culture media.

In some embodiments, the flow of media is driven by differences in Laplace pressure (see Figure 27). In Figure 11, there are two vertical stacks connected by a flow conduit 14, each stack is filled with culture medium 40, and interfaces between culture medium 40 and the immiscible liquid 2 are shaped like domes. Now, consider the case where stacks are of equal heights, and the holes 16 in cell holding sub-structures 10 are sufficiently narrow that interfacial forces dominate gravitational ones and the domes become shaped like the caps of spheres (see discussion of ‘balloons’ in Figure 12). Providing these caps have different radii of curvature, then differences in Laplace pressure can drive flow through the conduit 14, with flow direction being from the one with the smallest radius of curvature. Note that flow driven by hydrostatic pressure and by Laplace pressure is achieved without using a powered pump.

Specific Examples/Demonstrations

Specific examples and experimental demonstrations are discussed below with reference to Figures 12-26, and 28-40.

As mentioned above, in some embodiments different cell types may be grown in different cell holding sub-structures 10. The cell holding sub-structures 10 can thus act as microfluidic modules that can be used to build tissues (e.g. layers of different cells in different cell holding sub-structures 10). In the examples described below for demonstration purposes, the cells are grown in the holes 16 of cell holding sub-structures 10 of the type described in Figures 3-11. The cell holding structures 10 are provided mainly in the form of stainless steel washers (annular members), but in practice other shapes and compositions can be used. The cell holding sub-structures 10 can be formed from PTFE, hydrogels, or paper-based materials for example.

Figure 12 shows an image of a stack of stainless-steel washers acting as cell holding sub-structures. Cell culture medium (DMEM + 10% FBS) is provided in the middle of the stack (in the holes of the washers). The stack is provided in a 6 cm tissue culture dish filled with an immiscible fluorocarbon, FC40, acting as the cover liquid. The stack comprises five washers. Each washer is 1 mm thick, with an outer diameter, OD, equal to 10 mm, and an inner diameter, ID, equal to 5 mm. The stack is created simply by placing washers on top of each other in the dish in air. The washers are held in place by gravity without the need for any fixative or glue. Next, medium is added into the axial holes of the washers up to the top of the top washer. Some media seeps under the bottom washer and between other washers to their perimeters. It does not seep further, as it is held by interfacial forces acting at airmedia interfaces attached to stainless steel and polystyrene at the bottom, and to the stainless steel of two adjacent washers. FC40 is now added the dish to cover the stack. Finally, additional medium is pipetted through the FC40 to create the ‘balloon’ at the top. Perhaps counter-intuitively, media - which is less dense than FC40 - remains largely confined to the stack, and does not float above the FC40. This is because once media has wetted polystyrene and stainless steel, the added FC40 is unable to displace it, and interfacial forces confining the tower of media to the stainless steel in the stack above the footprint initially wetted with media. Although media seeps between washers to the edge of the stack, and between the bottom washer and the polystyrene substrate, media is confined to the stack solely by interfacial forces. No additional reagents or mechanisms (e.g., oils, surface treatments, gaskets, locking systems that keep the surface of adjacent washers closely together) are provided to prevent medium leaking from the stack. As FC40 is denser than media, buoyancy forces (i.e., the hydrostatic head of pressure) are driving media up (if more were present the ‘balloon’ of media at the top would detach and float off above the FC40). At this length scale, the ‘balloon’ has the shape of the cap of a sphere.

In some embodiments, to make a tissue or organ-on-a-chip/dish, each washer can contain one cell type, and different washers in the stack can contain different cell types. In tissues, cells adhere to each other, and to an extracellular matrix or scaffold. Therefore, each washer can, for example, contain a ‘scaffolding’ in the hole 16 that supports the cells. Many such scaffolds are available, including those known as ‘hydrogels’ and ‘aerogels’. With hydrogels, cells are usually dispersed by mixing them with the hydrogel in a liquid state, the liquid mixture is then dispensed into the appropriate place, and the liquid allowed to gel around the cells. Gelling can be achieved in many ways depending on the nature of the hydrogel involved, for example, by changing the temperature (as for agarose), creating chemical cross-links (e.g., using a cross-linking chemical, or by light when using a photo- activatable cross-linker). Here, liquid agarose plus cells would be poured into the hole 16 where gelation would occur so that the gel adheres to the sides of the hole. With an aerogel, the aerogel would be created in the hole 16 prior to adding cells. Aerogels can be made in many ways, and often involve removing water from a hydrogel, to leave a dry material with many interconnected cavities or pores. An example aerogel is one made with collagen and such aerogels are sold as substrates to make artificial skin. Collagen aerogels are often made by frothing collagen in acid, neutralizing to allow the collagen to set by forming cross-links, and freeze-drying to remove the water. Note that pores with diameters of a few hundred microns prove to be optimal for the growth of mammalian cells (e.g., those of human skin and bovine muscle), so the length scale of such pores is much the same as that over which diffusion and cell signalling work (above). In this case, an aerogel is created in the hole 16 so the aerogel adheres to the sides of the hole 16. Then, cell-containing media is dispensed on to the surface of the aerogel in the hole 16, the media and cells wick throughout the pores in the aerogel, the now-distributed cells adhere to the aerogel, and the cells grow on the underlying scaffold. The now-hydrated aerogel has become a hydrogel. Note that hydrated aerogel blocks can be used as the cell holding sub- structures 10 in Figures 1 and 2.

Once the different cell types have grown in the hydrogels in their respective washers in a stack (perhaps signalling to each other), the hydrogels in the stack can be pushed out onto a dish (e.g., using a plunger from a syringe if the holes 16 in the washers in the stack are circular). The result of this process is a stack of the different cell types (an ‘organ’). The order of the cell types can easily be changed by changing the stacking order of the washers. It is also possible to have mixtures of two or more cell types in one washer. Water in the cell growth environment is influenced by gravity (working through the high density of the cover liquid, FC40, and hydrostatic pressure). Interfacial forces (medium wetting the polystyrene of the dish and the stainless steel of the washers in this case) are important in determining how the water is retained in the cell growth environment defined by the washers.

The tissue pushed from the circular holes 16 of the washers has a circular footprint in this example. However, as described above, arranging for the holes 16 to have tessellating shapes, such as square or hexagonal, would allow the tissue units to tessellate and thereby build larger pieces of tissue, for example by side-by-side assembly to build artificial skin (see Figures 7 and 8 and the associated discussion above). As cells grow out from one cell holding sub-structure 10 to an abutting one in the tessellation, they will contact and adhere to cells attached to their neighbour and so create a bond between the two sub-structures 10.

If hydrogels (e.g. derived from aerogels) constitute the whole of cell holding substructures 10 in Figures 1 and 2, and - if the sub-structures have suitable shapes - they can be tessellated directly to build larger pieces of tissues without the need for prior pushing out of a holding washer.

It is important in embodiments involving washers for the hydrogel (or other suitable material) in the holes 16 of the washers (cell holding sub-structures) to stick effectively to the inner surface of the holes. It is therefore desirable to maximize a surface area of the washer that is in contact with the hydrogel, and minimize the weight of hydrogel so that it does not detach from the cell holding sub-structure during manipulation (e.g., stacking). Increasing a thickness of the washer, decreasing a diameter of the hole 16 in the middle, and/or introducing protrusions (such as ridges, ledges, threads, or surface roughening, as discussed above with reference to Figure 4) will help to ensure the hydrogel is reliably supported by the washer. When casting a hydrogel in a washer sitting in a dish, it is desirable to maximise sticking to the washer while minimising sticking to the dish so that the hydrogel does not detach from the cell holding sub-structure when it is lifted off the dish. The inventor has found that it is possible to cast various hydrogels as liquids (e.g., gelatin, agarose, alginate, collagen, cellulose-based mixtures) in the hole 16 of a washer while the washer is positioned in air on the surface of a suitable substrate. Suitable substrate surfaces include tissue-culture treated plastic, more hydrophobic bacteriological plastic, silanized glass, aluminium foil, and PTFE, with suitability being determined by the nature of the hydrogel. Whilst PTFE is attractive in principle as it is so unsticky, commercially available PTFE sheets are usually relatively rough so that cast hydrogels stick well to them. However, PTFE sheets can conveniently be made super-hydrophobic (and so less sticky) by applying an abrasive roughening process (sandpapering). Many other coatings that make sheets superhydrophobic are available (e.g., those using cellulose nanocrystal/SiCh).

In the above examples, hydrogels are cast on solid substrates. It is also possible to cast hydrogels on the smooth surface of a non-sticky liquid - the fluorocarbon FC40, for example. This is exemplified by the following experiment where -550 nl of an example hydrogel of 1% agarose in phosphate-buffered saline (plus red dye to improve visibility) is cast on liquid FC40 (which is denser than water) into the 7 holes of 7 Ml hexagonal nuts, each initially incorporated into 7 stacks in a 6 cm dish. Each nut had an inner diameter nominally equal to 1 mm with a thread, a maximal outer diameter of 2.71 mm, and a height of about 0.7 mm. Each stack contained a PTFE washer in the bottom layer, and the nut plus a metal washer in the second layer. The metal washer acts solely as a weight to prevent the PTFE washer on which it sits from floating away when dense FC40 is added later. The centres of the nut, PTFE washer, and metal washer were co-axially aligned. The PFTE washer had an inner diameter of 2.5 mm (slightly less than the maximal outer diameter of the nut), and the nut sat over the hole in the PTFE washer. The metal washer in the second layer had an internal diameter of 5 mm (slightly larger than the maximal outer diameter of the nut), and sat on the PTFE washer around the nut. FC40 was added to the dish, up to the top of the PTFE washer, and level with the bottom of the nut. Then, molten agarose was cast in the hole in the nut and onto the FC40, and allowed to set as it cooled. Each of the 7 nuts (with adherent agarose in the hole) were removed, and transferred through FC40 and air into a new 7-layer stack in the dish under FC40. This shows that a typical hydrogel can be cast on a non-sticky liquid surface and then removed from that surface, and that hydrogels can adhere even to the stainless steel in nuts as stacks are created and rearranged. Note that hydrogels stick to paper-based ‘washers’ (see below) even more tightly than to stainless steel ones.

In the above examples, the hydrogel was cast in the hole in a washer. More generally, whenever a hydrogel (or what will become an aerogel) is cast on a flat surface prior to use as a cell-holding sub-structure, it is desirable to minimise sticking of the hydrogel to the flat surface so that the set hydrogel can be removed easily. Then, hydrogels not in the holes in washers can be cast on non-sticky solids or liquids in the ways described above.

During cell culturing, it is necessary to feed cells in the stack as they grow. Figure 13 depicts an example stack comprising three washers (what looks at first glance like a fourth at the very bottom is a reflection). The actual bottom washer has three 1 mm spacers above it so it is possible to see medium (and two of the spacers, as one is hidden behind a second) in the gap. This stack was built in air, and then media pipetted into the hole in the middle of the bottom washer up to the height of the bottom washer; then, FC40 was added to roughly the same height. Next, media was added up to the top of the second layer (so wetting the spacers) before FC40 was added up to the same height (i.e., the top of the spacers). This process of progressively adding media before FC40 was repeated for the final two layers, before finally the stack was covered with FC40. This progressive filling of the system is performed to avoid unwanted displacement of media by FC40 during the filling, and to ensure that media wets stainless steel in the next layer before FC40, so that media extends to the edge of the stack. Note that if FC40 wets spacers before adding media, the additional media can fail to displace FC40 from the stainless steel, and this gives a narrower central column of media in the stack. The gap in the layer with spacers is wide enough to insert a PTFE tube through the FC40:media interface and thereby pump medium into the middle to feed cells in the stack. Note also that when a PTFE tube is inserted through the FC40:media interface, the interface automatically seals around the inserted tube so that no media is lost from the stack. Similarly, the FC40:media interface automatically reforms when the tube is withdrawn.

Figures 14-17 depict an alternative approach in which feeding occurs via a lateral flow over a bridge between different stacks. Figure 14 shows two stacks in an empty dish in air. The left stack has 5 washers. The right stack has 6 washers. A bridge acting as a flow conduit is provided between the two stacks. The bridge in this example comprises a piece of Whatman No 1 filter paper with a nominal thickness of 180 microns. Figure 15 shows the arrangement after cell culture medium has been added to the stacks (in the holes of the washers) and the stacks have been overlaid with FC40 as a cover liquid. Media has wicked across the bridge. Figure 16 shows the arrangement after further medium is added to the stacks. When the arrangement of Figure 16 is left alone, the stack having the greatest height of medium, which is the stack on the right, pulls medium across the bridge from the stack that has the least medium, due to hydrostatic pressure, leading to the situation shown in Figure 17. The situation is similar to that discussed above with reference to Figure 11. The experiment shows that one can add modules together to form stacks and feed media both vertically and laterally without using a powered pump. Note that flow can also occur downwards when using an overlay that is less dense than water.

Figures 18-20 depict an alternative sequence in which the stacks are fed from below. As shown in Figure 18, two pairs of washers are provided in this example. Each pair of washers forms a separate stack. Both pairs of washers are provided on a strip of Whatman filter paper (which will act as a flow conduit). Any other kind of flow conduit could be used instead. In a subsequent step, the filter paper was wetted with cell culture media, FC40 was added over the top as a cover liquid, and then more media was added into each stack to fill each stack. The resulting arrangement is shown in Figure 19. Media in the two stacks were now demonstrated to be fluidically connected. Media was added to the left-hand one, bubbles made during this process were detached. Transfer of media to the right-hand stack was then observed, as shown in the sequence of images of Figure 20 (which follow progress of the experiment as a function of time from top to bottom).

In the examples above, the washers were made of stainless steel, to which biocompatible hydrogels and scaffolds stick with difficulty. Hydrogels will stick to other materials better. Hydrogels stick to paper very well, for example. Figure 21 shows a stack in a 6 cm dish with medium in the middle, and an FC40 overlay. One layer in this stack is made of a paper tube constructed from 2 layers of Whatman No 1 filter paper (this paper has a nominal thickness of 180 microns). The tube is ~2 mm high and ~8 mm across. This stack has (from bottom up): washer (1 mm thick, outside diameter 10 mm, inside diameter 5 mm)

3x 1 mm high tubular metal spacers (diameter 1 mm) creating the gap washer (1 mm thick)

~2 mm high paper tube (diameter ~8 mm) - this would contain, for example, a scaffold + cells washer (1 mm thick).

Medium was added in the middle to the height of the bottom washer, then FC40 to roughly the same height. This process was repeated for each layer, before finally covering the stack with FC40 as a cover liquid. This progressive filling of the system is performed to avoid unwanted displacement of media by FC40 during the filling.

In Figure 21, the cover liquid (FC40) retains media within the cell growth environment by interfacial tension at FC40:media interfaces across a plurality of openings with a wide range of sizes. For example, the widest opening (with the ‘balloon’ at the top) has a diameter of 5 mm. The next widest (~1 mm high) are at the sides of the layer with the spacers (i.e., between the two bottom washers); these openings are still large enough to allow insertion of a tube into the gap so media can be pumped in to flow automatically up into the cap due to hydrostatic pressure. A plurality of the smallest openings are on the outer face of the tube of filter paper, and have dimensions down to a few microns. When a stack like this is over-filled with media, it has never been observed that media is lost through the smallest openings. This is to be expected, as the pressure difference across interfaces at the openings is given by the Laplace equation, and inversely depends on the principal radii of curvature, with the ones at the smallest openings being the smallest.

Figures 12-21 provide examples with stainless-steel washers in stacks. The use of stainless steel serves two main purposes: it is easy to pick up by hand or with forceps when building these exemplary stacks, and it acts as a weight to stabilize the stack both in air and when under dense FC40. Whilst stainless steel can be used in a cell holding sub-structure 10 (as in Figure 3), it can be replaced by any scaffold suitable for cell growth that is strong enough to retain its shape when assembling stacks. The use of paper instead of stainless steel has many attractions.

We will use the term ‘paper’ for any sheet of aerogel, sponge, hydrogel, or scaffold that can be cut easily (e.g., with scissors or a scalpel) into a wanted 2D shape. It includes sheets made with cellulose-based materials from bacteria and plants (e.g., trees, sugarcane, cotton, hemp), edible starch-based wafer papers and rice papers, and filter papers used by chemists to remove particulates like Whatman No 1 papers. The chemistry of many such papers is well developed, and their surfaces can be modified in almost any wanted fashion (e.g., by covalent attachment of the ‘RGD’ tripeptide consisting of arginine, glycine and aspartate that promotes cell attachment). Additionally, such papers have been widely used as scaffolds when culturing a variety of cell types. Consequently, use of cell holding sub-structures 10 made wholly of ‘paper’ are attractive modules when building (layer by layer and/or side-by-side) an ‘organ’ containing different ‘tissues’ with almost any chosen 3D shape (e.g., a portion of skin like an ear lobe, heart, kidney, or a T- bone steak).

In the example of Figure 21, one layer contains a Whatman No 1 filter paper rolled into a ring. Figures 22 and 23 depict an alternative arrangement in which four square filter papers are used to build a stack so that stack height is easily varied. Each filter paper in this example was -180 microns thick and 10 mm square with a 3 mm square hole in the middle. The filter papers are sandwiched between two washers in a stack (the top one acts as a weight). Figure 24 depicts the arrangement of Figures 22 and 23 after adding media and overlaying with FC40 as a cover liquid. As paper is available in wide range of thicknesses (e.g., Rizla super thin silver cigarette rolling paper is 20 microns thick, and Whatman filter papers Numbers 50, 1, 2, 4, 3 are 100, 180, 190, 210, and 390 microns thick, respectively), layer thickness can be varied easily.

As mentioned above with reference to Figure 11, in some embodiments the hydrostatic head can be used to drive a pulsatile flow of the cell culture media. For example, a dish containing two stacks connected by a bridge and covered with a cover liquid can be mechanically rocked on a rocker. Alternatively, a long thin rectangular ‘washer’ with a rectangular hole containing hydrogel could be rocked. Alternatively, pulsatile flow could be obtained as follows. A stack of two long thin rectangular washers (each with a hydrogel) with spacers in between is provided. The middle layer is filled with medium, and has open walls along the length of the rectangle. Cells, such as endothelial cells, could be plated in the middle so that the cells line the hydrogel ceiling and floor. The cells are then grown so that the cells fill the FC40/medium wall. The arrangement is then rocked to provide a pulsatile flow, thereby providing a primitive ‘vasculature’.

Figures 25 and 26 depict two hydrogels in a stack built as follows (the two hydrogels could contain the same or different cell types):

1. Put stainless steel washer (e.g. OD = 10 mm, ID = 5 mm) in 6 cm dish, fill hole with 50 pl DMEM (this is a reservoir of media), add 1 ml FC40 (which does not cover the washer).

2. Add a tube made of Whatman filter paper No 1 containing 1% agarose in phosphate buffered saline (PBS) with red dye (this was cast in air on a hydrophobic dish made with bacteriological plastic), add 2 ml FC40 (to bring the top of the FC40 almost up to the top of the bottom washer), and put 25 pl DMEM directly on top of the agarose.

3. Add second tube of Whatman filter paper with agarose, put 2nd stainless steel washer on top (which acts as a weight), add 4 ml FC40, fill hole in washer with 50 pl DMEM, top up with 12 ml FC40 (to completely cover the stack).

This approach provides two reservoirs of cell culture media (in the holes of the washers at the top and bottom), and two abutting hydrogel discs in the middle of the stack (each contained in a filter-paper tube). The aqueous phases in the two discs are in continuous liquid communication with each other. Media can easily be refreshed in the top reservoir by pipetting. If it is desirable to refresh media from the bottom, it is possible to include extra layers (e.g., using spacers to provide an open side so a tube can deliver media to an internal reservoir as in Figure 13, or through a Whatman filter paper on the bottom).

Figure 27 depicts an example cell holding structure comprising a stack of two cell holding sub-structures 10 in the form of washers provided on a bottom inner surface of a container 6 (polystyrene dish). Media is added to the centre of the stack (in the holes 16 of the washers). The container 6 is filled with FC40 acting as the cover liquid 2. Air is provided in the region 4 above the interface 5 of the cover liquid 2. The plan and side views in Figure 27 illustrate the result. Media is shown bulging out at P3 and P4 (i.e., the media has a convex face). This is the case in air. However, depending on the height of the FC40 overlay and the wetting sequence (see discussion of Figure 13 for the effects on the fluidic structure of the wetting sequence), media can bulge inwards (i.e., have a concave face).

In this example, media has wicked from the centre of the stack between the container 6 and the bottom washer, and between the two washers - the media wet the steel and polystyrene before the FC40. However, interfacial forces limit further wicking either along the container surface 6, sideways between the washers, or out at the top. Media is ‘pinned’ to the metal or polystyrene at the edge of the media:FC40 interfaces. Successful leaking would increase the wetted surface of the container 6, or lead to media ballooning up from between the two washers or from the top (when it would break free to float up above the denser FC40). The latter is the usual case when more media is added, but ballooning up from the side can occur if the distance between washers at P3 is increased (e.g., if longer spacers than those shown in Figure 13 were used between the two washers).

Prevention of leaking is desirable. The main forces to be considered that prevent leakage are interfacial and hydrostatic forces (FC40 being denser than water). The difference in pressure across the media:FC40 interfaces at points P2, P3, and P4 is governed by Laplace’s law, and so on the radii of curvature (i.e., Reap, Rside, and Rbottom). Consequently, the Laplace pressure difference across the interface at P3 is greater than at P2. The fluidic structure is stable as long as the net buoyancy force acting in/around P2, P3, and P4 - which drives media up - is less than counter-balancing hydrostatic + Laplace pressures.

Imagine pumping in more media at P5 (e.g., through a pipet tip introduced from above). First, interfaces at P2, P3, and P4 bulge out to accommodate the additional volume, until pinning lines are eventually broken and media balloons up to float to the surface. In principle, loss of pinning can occur at any of the interfaces around P2, P3, and P4. In practice, ballooning up from P2 is generally seen. Ballooning up from P4 has not been observed unless the stack is sitting on a piece of filter paper connected to another stack. Ballooning up from P3 has not been observed unless there are tall spacers between the two washers (e.g., 4 mm high spacers used in a stack like that in Figure 13). The point at which bubbling up occurs depends on the interplay between buoyancy forces and Laplace pressures curvatures (and so on the radii, Reap, Rside, and Rbottom). In Figure 21, the curvatures in the plenitude of openings in the sides of the tube of filter paper are high, so when such a stack is over-filled, media has never been observed to be lost through such openings in the filter paper, or from openings in the sides of hydrogels.

It is possible to determine the pressure at P0, for example, in two ways: by considering pressures along the line from air straight down through P2 (route 1, which includes contributions due to the hydrostatic head h2 of the cover liquid 2, plus that due to the Laplace pressure at the interface with radius Reap, plus that due to the hydrostatic head h3-h2 of the media), or following a dog-leg straight down to P3 and then sideways (route 2, which includes contributions due to the hydrostatic head h3 of the cover liquid 2, plus that due to the Laplace pressure at the interface with radius Rside). These two estimates must give the same answer, so one can relate curvatures of interfaces at P2 and P3.

Fluid flow through a porous medium like a hydrogel is governed by Darcy’s Law. The flow depends on the pressure difference and flow resistance of the porous medium, and is analogous to Ohm’s law (with flow being equivalent to current, the pressure difference to voltage difference, and flow resistance to electrical resistance). The generic flow resistance term in Darcy’s law includes contributions from viscosity, the length of porous medium traversed, medium permeability, and cross-sectional area. When feeding cells growing in cell holding sub-structures 10, it is often desirable to have sufficient flow through the sub-structures 10 to provide nutrients and remove waste to maintain viability. In general, flows take the path of least resistance (e.g., between/around sub-structures 10 over their surfaces, and/or through sub-structures 10 that have the least flow resistance). Consequently, flow patterns through a set of tessellated sub-structures 10 with different flow resistances (e.g., through stacks where layers contain cells of different types and densities growing in different hydrogels or matrices) that may be packed together more or less closely are complex. At a simple level, such flows are analogous to electrical flows through different electrical resistances arranged in series and/or in parallel. As with electrical circuits, if there is a pressure difference across the system (equivalent to a potential drop), then some flow will occur through all sub-structures 10 irrespective of their flow resistance.

Some cell holding sub-structures 10 may have such high flow resistances that flow through them is too low to maintain cell viability. However, providing cells lie within diffusional reach (i.e., -200 microns) of a region with a high enough flow (which might be over an external surface, in the space between two adjacent sub-structures 10, or through an adjacent sub-structure 10 with a low flow resistance), sufficient nutrients can still be delivered and waste removed to maintain viability.

Figures 28-40 depict an experiment showing flow through a (cell-free) cell holding structure 8 containing two different types of cell holding sub-structure 10 labelled 10A and 10B. The experiment also illustrates how easy it is to assemble and disassemble a cell holding structure 8 and establish flows through it. Figure 28 is a schematic side sectional view illustrating the experimental configuration. Figure 29 is a top view prior to starting flow (5 minutes before). Figure 30 is a side view 30 minutes after flow has started.

The two different types of cell holding sub-structure 10A and 10B have different flow resistances relative to each other. The first type of sub-structures 10A have relative high resistance and are provided in the form of gelled blocks of 3% agarose in phosphate- buffered saline, PBS. The first type of sub-structures 10A may be referred to as agarose blocks (or simply “blocks”). In the example shown, the agarose blocks are provided in two stacks in zones B and C. Individual blocks are labelled Bl, B2, Cl and C2. The second type of sub-structures 10B have relatively low resistance and are provided in the form of Whatman No 1 filter paper. Washers 34 and nuts 36 are provided as weights and/or spacers. Another spacer layer 38 separates the stack of sub-structures 10A-10B in zones B and C from overlying nuts 36. The cell holding structure 8 is provided in a 6 cm polystyrene dish containing FC40 as a cover liquid 2 to a level indicated by the broken horizontal line. The outside environment 4 is air. Liquids can be added to the lowermost sub-structure 10B via a dispensing needle 32.

In this example, the 6 cm polystyrene dish is not tissue-culture treated to minimise aqueous flows around the lowermost sub-structure 10B (filter paper), which sits on the bottom of the dish. A row of four stacks (forming respective zones A-D) are built in the dish. Stacks in zones A and D form input and output reservoirs respectively. Stacks in zones B and C contain the sub-structures 10A-10B (porous layers of agarose blocks and filter paper). Some or all of the sub-structures 10A-10B could contain cells.

The structure may be built as follows.

1. A rectangular piece of Whatman No 1 filter paper (28 x 8 mm) is placed on the bottom of the dish, wetted with 25 pl PBS, and most air bubbles removed by pressing on the paper. This paper will constitute the bottom layer of stacks A-C and is one of the substructures 10B. It may be referred to as lower sub-structure 10B. Pre-wetting the paper with PBS ensures the paper remains stuck to the dish (held by interfacial forces) when the cover liquid 2 (in this case FC40) is overlaid.

2. Stack A (the input reservoir) is completed by placing the center of a stainless steel washer 34 (OD 10 mm, ID 5 mm, thickness 1 mm) on the midline of lower sub-structure 10B, so the left-hand edge of the washer 34 extends ~2.5 mm beyond the left-hand end of the lower sub-structure 10B. The washer 34 also serves as a weight that provides an additional way of holding the lower sub-structure 10B on the bottom of the dish when FC40 is added. This arrangement creates an overhang of washer 34 over lower substructure 10B that serves to prevent PBS which is added later from seeping up the outside edges of the washer 34 when FC40 is overlaid.

3. The second layer of the stacks in zones B and C each consists of one square block of (gelled) 3% low-melting agarose in PBS (footprint 8 x 8 mm, height 3 mm). Each block is one of the sub-structures 10 A. We will call these blocks Bl and Cl. They are stored in PBS prior to use and placed while still wet with PBS on lower sub-structure 10B. The centers of both Bl and Cl are on the center-line of lower sub-structure 10B, and the righthand edge of Cl extends ~2 mm over the right-hand edge of lower sub-structure 10B. The right-hand edge of Bl abuts the left-hand of edge of Cl to provide aqueous continuity between the two. This placement of blocks on lower sub-structure 10B ensures there is a space between the washer 34 in the stack of zone A and Bl. As each square block is wider than lower sub-structure 10B, Bl and Cl overhang the sides of lower sub-structure 10B, and Cl also overhangs the right-hand end of lower sub-structure 10B. During operation, some left-to-right flow will occur through the portion of the lower sub-structure 10B under each block. Some flow may also occur on each side of the lower sub-structure 10B (i.e., under the overhang), and the overhang will minimise flow up the sides of blocks Bl and Cl when FC40 is overlaid. Some flow will also occur through each block, although this will be less than through the lower sub-structure 10B, as paper has a lower flow resistance than 3% agarose.

4. The third layer of the stack in zone B consists of another agarose block (a sub-structure 10A), referred to as B2. The third layer of the stack in zone C consists of another agarose block (a sub-structure 10 A), referred to as C2. Blocks B2 and C2 are identical to blocks Bl and Cl, and B2 is placed on Bl and C2 is placed on Cl. Blocks Bl and B2 are gently pushed sideways on to blocks Cl and C2 to ensure lateral aqueous continuity between blocks in stacks B and C. As all blocks are wet with PBS, aqueous continuity is established with neighbouring blocks when they are pushed together. Additional layers will be added later to the stacks in zones B and C.

5. The stack in zone D (the output reservoir) is built on the dish several millimeters to the right of the half-built stack in zone C. A stainless-steel M6 nut 36 (width across flats, S = 10 mm; thickness, M = 5.2 mm) is placed on the dish, and then a stainless-steel washer 34 (OD 10 mm, ID 5 mm, thickness 1 mm) is placed co-axially on top of the nut 36.

6. A rectangle of Whatman No 1 filter paper acting as a sub-structure 10B (which may be referred to as upper sub-structure 10B) - which is pre-wetted with PBS - is now added along the top of B2, C2 and the washer 34 in stack D. The left-hand edge of upper substructure 10B is positioned ~ 2 mm to the right of the left-hand edge of block B2. Upper sub-structure 10B has the same dimensions as lower sub-structure 10B, but additionally has a rectangular hole (4 x 3 mm) close to its right-hand end. The right-hand edge of this hole is 3 mm away from the right-hand end of the upper sub-structure 10B; consequently this hole lies over the holes in the washer 34 and nut 36 below.

7. PBS (100 pl) is now pipetted from above directly through the co-axially arranged holes in the growing stack in zone D on to the bottom of the dish. This PBS wets both dish and stainless steel nut 36 and is held by interfacial tension within the nascent stack in zone D.

8. FC40 (4 ml) is now added to the dish; it covers the washer 34 in the stack in zone A, and minimises seepage of PBS out of the nascent stack in zone D. Interfacial tension plus the effects of gravity hold the lower sub-structure 10B and the four agarose blocks to the bottom of the dish.

9. A second M6 nut 36 is added to complete the stack in zone D. This nut 36 will prevent the right-hand end of the upper sub-structure 10B from detaching from the stack in zone D and floating away when much of the arrangement is overlaid later with FC40.

10. A solid rectangle of PTFE (25 x 13 x 1 mm) acting as a spacer layer 38 is placed over the upper sub-structure 10B above blocks B2 and C2. It provides a roof over most of the upper sub-structure 10B, and acts as a spacer layer 38 between the upper sub-structure 10B and two M6 nuts 36 that will be added later. This rectangle is gently pressed down on to nascent stacks in zones B and C to ensure there is vertical aqueous continuity between the upper and lower sub-structures 10B and the agarose blocks that lie under the rectangle. This PTFE rectangle is longer and wider than the underlying agarose blocks, and it also overhangs the left-hand of the upper sub-structure 10B; this overhang will eliminate any upward flow of the aqueous phase around the edges of the rectangle when FC40 is overlaid later. The stacks in zones B and C are now completed by the addition of an M6 nut 36 to each stack on top of the PTFE rectangle. These nuts will prevent the underlying upper and lower sub-structures 10B, agarose blocks, and PTFE rectangle from floating away when FC40 is added later. All stacks are now complete.

11. A further 5 ml FC40 is added to fill the dish up to a level that is about half way up block B2, 100 pl is pipetted into the PBS already in stack D, and the dish is completely filled with FC40. Pressures in the system now equilibrate over the next ~15 min. Note that unknown amounts of PBS are added to the system when pre-wetted filter papers and agarose blocks are added during stack building, so sometimes a ‘balloon’ of PBS can be seen at the top of the stack in zone D, and sometimes this balloon can shrink or expand during equilibration. It is convenient to add/remove PBS to the balloon by pipetting through FC40 so the top of the balloon can just be seen from the side above the top of the top-most nut in the stack in zone D; then, when PBS +/- dye is pumped into the stack in zone A, an increase in size of this balloon allows easy monitoring of successful transfer of the aqueous phase through the system from left to right and into the stack in zone D.

12. The photo in Figure 29 labelled ‘-5 min’ gives a top view of the filled dish at this stage. Layers in stacks are as follows (from bottom to top). In the stack in zone A: lower sub-structure 10B, and steel washer 34. In the stack in zone B: lower sub-structure 10B, agarose block Bl, agarose block B2, upper sub-structure 10B, spacer layer 38 (PTFE rectangle), and steel nut 36. In the stack in zone C: lower sub-structure 10B, agarose block Cl, agarose block C2, upper sub-structure 10B, spacer layer 38 (PTFE rectangle), and steel nut 36. In the stack in zone D: steel nut 36, steel washer 34, upper sub-structure 10B, and steel nut 36. Note that the plastic dish is hydrophobic, PTFE is fluorophilic and hydrophobic, and stainless steel is hydrophilic. The arrangement exploits these properties to minimise seepage of PBS under, over, and around the lower and upper sub-structures 10B and agarose blocks, and to contain PBS in input and output reservoirs. While a syringe pump is used to introduce PBS into the stack in zone A, it will be differences in hydrostatic pressure that are the main drivers of flow from left to right and upwards, with Laplace pressures in the ‘balloon’ in the stack in zone D playing only a minor role. However, Laplace pressure plays a major role in retaining the aqueous phase within and between the lower and upper sub-structures 10B and agarose blocks.

13. The tip of a stainless-steel dispensing needle 32 is now lowered down through the hole in the washer 34 in the stack in zone A until it touches the wetted lower sub-structure 10B. This needle 32 is fitted to a ‘holder’ that rests on top of the dish. The needle 32 is also filled with PBS plus blue dye (0.4 mg/ml resazurin), and connected via a Teflon tube to a 1 ml syringe housed in a syringe pump. Once the pump starts, PBS plus blue dye is delivered at 50 pl/h to the lower sub-structure 10B in the stack in zone A. Thereafter, PBS plus dye will flow through the structure to the balloon at the top of the stack in zone D. The photo in Figure 30 illustrates a side-view of the set-up 30 min after starting the pump; close inspection reveals that some blue dye has already passed along the lower sub-structure 10B and is entering block Bl (marked by the arrow labelled ‘front’). Note that the positive times given in photos shown in Figures 30 onwards refer to the time that the pump has been in operation and not actual time elapsed, as times taken later to dismantle stacks, photograph agarose blocks, reassemble stacks, and refill the pump are all excluded (in combination these processes take only ~15 min whenever a stack is dismantled or assembled).

Figures 31-40 illustrate flows through the cell holding structure 8 described above with reference to Figures 28-30, and how easy it is to dismantle and reassemble the structure and re-establish flows. It was seen in the discussion with reference to Figures 28- 30 that the dye front was just entering agarose block Bl 30 min after starting the pump.

Two hours after starting the pump, the balloon on top of the stack in zone D (see arrow in the top panel of Figure 31) has been seen to grow as water is transferred through the system. Flow is largely due to a difference in hydrostatic pressure, as the head of dense FC40 over the stack in zone A is greater than that over the stack in zone D. As the experiment runs, PBS in the balloon is removed manually by pipetting, or - during the night - automatically, as PBS floats up (and over) the FC40 overlay to contact either the edge of the nut 36 on top of the stack in zone C (which sticks up above the fluorocarbomair interface), or the top edge of the dish. Three hours after starting the pump (middle panel in Figure 31), dye has entered the bottom left-hand comer of block Bl, and the front is following a path of least resistance in the gap between blocks Bl and Cl. Seven hours after starting the pump (bottom panel in Figure 31), dye has filled all four agarose blocks and the lower and upper sub-structures 10B. During this initial phase, air bubbles spontaneously emerge from solution and/or the upper sub-structure 10B to float up to the FC40:air interface to be lost to the system (the arrow in the bottom panel of Figure 31 indicates one such bubble temporarily trapped under the PTFE rectangle); this occurs without disrupting flow. Note that trapped air bubbles often cause catastrophic failures of conventional microfluidic devices, but have no effect here.

The assembly is now dismantled: the pump is stopped, the dispensing needle 32 raised, the stack in zone D emptied of PBS (now containing some dye) by pipetting, the dish emptied of FC40, and stacks dismantled (from the top down). Now, the four agarose blocks and upper and lower sub-structures 10B are passed through FC40, and then FC40 drained from their surfaces; this ‘rinses’ away excess PBS and dye from their surfaces. Next, the four blocks are placed on a dish and photographed (Figure 32). Block Bl contains the most blue dye, which on this occasion is concentrated towards the bottom left. This is probably because dye flowed more rapidly along the front of lower sub-structure 10B compared to the back (perhaps because more air bubbles were initially present at the back where they reduced flow). Block B2 contains less dye than Bl, while C2 (which was most distant from the input) contains the least. A pressure difference across blocks coupled to diffusion has contributed to the entry and exit of dye into, between, and out of blocks.

All four agarose blocks plus the lower and upper sub-structures 10B are now rinsed briefly in PBS, reassembled as before into four stacks, and PBS plus blue dye pumped into the stack in zone A for another 16 h. The photo in Figure 33 (now t = 23 h) illustrates the structure after removing the dispensing needle 32. All four agarose blocks are densely blue, and dye has reached the stack in zone D. The structure is now dismantled as before, and isolated agarose blocks rephotographed (Figure 34); each block now contains more blue dye than at 7 h. The structure is now reassembled, and blue dye flushed from it by pumping PBS without dye (at 50 pl/h) into the stack in zone A for 7 h. The photograph in Figure 35 (t = 30 h) shows that the agarose blocks contain less dye than at t = 23 h. After dismantling the structure, a photograph of isolated blocks (Figure 36) confirms they contain less dye than at 23 h.

The structure is now reassembled, more PBS without dye pumped into the stack in zone A for 16 h, and another photograph taken (Figure 37, at t = 46 h); agarose blocks contain less dye than at 30 h. After dismantling the structure, a photograph of isolated blocks (Figure 38) confirms they contain less dye than at 30 h.

The structure is now reassembled, more PBS without dye pumped onto the stack in zone A at a five-fold higher rate (i.e., at 250 pl/h) for 3.5 h, and another photograph taken (Figure 39, at t = 49.5 h); agarose blocks contain less dye than at 46 h. After dismantling the structure, a photograph of isolated blocks (Figure 40) confirms they contain less dye than at 46 h.

The results illustrated in Figures 28-40 illustrate the ease of assembly of cell holding sub-structures 10 into a larger cell holding structure 8, and how flows through the system can be started and stopped. When creating tissues, organs, and food, PBS will be replaced by an appropriate cell-growth medium, and some or all of the sub-structures (e.g. filter papers and agarose blocks) will contain cells of appropriate types.

Clonal growth of HEKs in modules containing Geltrex

Modules (cell holding sub-structures) can be made of any appropriate biomatrix that can be picked up and stacked. Materials too fragile to be handled alone can be cast in more solid scaffolds or meshes; for example, we have filled stainless-steel and paper washers with agarose, collagen, Matrigel™, Geltrex™, alginate, and cellulose (both nano- fibrillar and nano-crystalline). Use of cellulose, starch, and fibroin-based scaffolds are particularly attractive as they are robust, biofriendly and biodegradable; they are also available as sheets with thicknesses down to ~20 pm that can easily be cut into any desired 2D shape. We now illustrate clonal growth of human embronic kidney 293 cells (HEKs) in Geltrex hydrogels filling cavities in meshes made of bio-inert nylon - chosen because it is widely available in sizes spanning our desired length scales and cell growth can easily be monitored easily in individual mesh chambers it in real time using a standard microscope. Meshes are dipped successively in ice-cold Geltrex containing HEKs so each mesh chamber fills with liquid hydrogel and a few cells, and gels set by incubation at 37°C (for 10 s in FC40); then, meshes are transferred to medium where they float freely. After culture overnight, most chambers in the mesh contain a few single cells (Figure 41 (i)) that grow into clones over the next two weeks (Figure 41 (ii)). Other meshes are stacked, and cells fed by pipetting medium on to a filter-paper at the base (Figure 41 (iii), cartoon). On day 7, the stack is disassembled, and central chambers from the center of the central mesh in the stack are imaged; after reassembling the stack, feeding continues for another 7 d, the stack is disassembled again, and similar central chambers re-imaged. In general, clones in stacks appear marginally larger than controls (Figure 41 (iii), images), but it is difficult to compare volumes precisely due to differences in procedures. For example, some colonies grow out of gels in free-floating meshes to attach to dishes, while those in stacks grow on surfaces between stacks and/or are tom from their parent colony during unstacking. Note that flow through central regions is easily increased by piercing an assembled stack several times with a needle, and that disassembly is equivalent to - and easier than - the sectioning required to image cells at the center of organoids grown conventionally. These results show that HEKs clone normally in stacks even when at least 500 pm from stack sides and 850 pm from the top or bottom - distances at the upper end of the diffusional range (these are under-estimates as hydrogel coats the outsides of individual meshes to increase thickness).

Claims

1. A method of culturing cells, comprising: providing a cell holding structure having a plurality of openings, wherein the cell holding structure contains an aqueous cell growth environment, and the cell growth environment is submerged in a cover liquid that is immiscible with water and retains water within the cell growth environment by interfacial tension at interfaces between the water and the cover liquid across at least a subset of the openings; providing living cells in the cell growth environment; and culturing the cells in the cell growth environment by providing a flow of cell culture media into and out of the cell growth environment through at least one of the openings while the cell growth environment is submerged and confined by the cover liquid.

2. The method of claim 1, wherein the cell holding structure comprises a plurality of detachable cell holding sub-structures, each cell holding sub-structure comprising a plurality of openings and containing a respective portion of the aqueous cell growth environment.

3. The method of claim 2, wherein the providing of the cell holding structure comprises moving initially separated cell holding sub-structures into contact with each other.

4. The method of claim 3, wherein living cells are provided in some or all of the cell holding sub-structures before the cell holding sub-structures are moved into contact with each other.

5. The method of claim 4, wherein the living cells are cultured in the cell holding substructures before the cell holding sub-structures are moved into contact with each other, the culturing being performed by providing a flow of cell culture media, driven by a pressure gradient, into and out of each cell holding sub-structure through at least one of the

35 openings in each cell holding sub-structure while the cell holding sub-structure is submerged by the cover liquid.

6. The method of any of claims 3-5, wherein water is held in the cell holding substructures by interfacial tension at interfaces between the water and surrounding medium during the moving of the cell holding sub-structures into contact with each other.

7. The method of claim 6, wherein each of at least a subset of the interfaces between the water and surrounding medium is replaced by a continuous aqueous liquid connection between the respective cell holding sub-structures when the cell holding sub-structures contact each other.

8. The method of any of claims 3-7, wherein the cell holding sub-structures are moved into contact with each other while the cell holding sub-structures are submerged in the cover liquid.

9. The method of any of claims 2-8, wherein all of the respective portions of the aqueous cell growth environment in the different cell holding sub-structures are in continuous aqueous liquid communication with each other via at least a subset of the openings in the cell holding sub-structures.

10. The method of any of claims 2-9, wherein each of one or more of the cell holding sub-structures comprises openings in an upper surface and a lower surface to allow vertical stacking of the cell holding sub-structures to form the cell holding structure.

11. The method of any of claims 2-10, wherein each of one or more of the cell holding sub-structures comprises openings in a lateral surface to allow horizontal stacking of the cell holding sub-structures to form the cell holding structure.

12. The method of any of claims 2-11, wherein two or more of the cell holding substructures contact each other via respective planar surfaces that each contain plural

36 openings.

13. The method of any of claims 2-12, wherein two or more of the cell holding substructures contain biological cells of different types relative to each other so that the culturing of cells produces a structure of cells of different type in different regions.

14. The method of any of claims 2-13, wherein each cell holding sub-structure is a porous body and at least a subset of the openings are formed by pores in the porous body.

15. The method of claim 14, wherein the porous body comprises a matrix, scaffold, polymer or hydrogel formed from units of sugar, carbohydrate, amino-acid or hydrocarbon residues.

16. The method of any of claims 2-15, wherein each cell holding sub-structure comprises a substantially planar member having a hole extending through the plane of the planar member, the hole defining a respective opening on each side of the planar member.

17. The method of claim 16, wherein the holes of two or more of the cell holding substructures are aligned with each other vertically to form at least a portion of the aqueous cell growth environment.

18. The method of claim 16 or 17, wherein a radially inward facing wall defining the hole of each of one or more of the planar members comprises one or more protrusions.

19. The method of any preceding claim, wherein the cell culture media is directed into the cell growth environment along a microfluidic conduit, along a tube, or along porous material.

20. The method of any preceding claim, wherein the flow of cell culture media is driven by a pressure gradient.

21. The method of any preceding claim, wherein the pressure gradient is applied predominantly using hydrostatic pressure.

22. The method of any preceding claim, wherein the flow of cell culture media is varied to provide a pulsatile flow.

23. The method of claim 22, wherein the pulsatile flow is provided by mechanically rocking the cell holding structure and the cover liquid.

24. The method of any preceding claim, wherein the cover liquid is denser than water.