Bench philosophy: Closer to Real Life
3D Cell Culture
by Steven Buckingham, Labtimes 01/2013
Just like in the world of cinema, cell culture is moving from 2D to 3D.
Once, we were happy to grow cells as flat sheets; it suited us and it simplified things; it kept things nicely under control. But all along we were missing something and eventually we recognised the obvious: cells don’t live in a 2D world. Cells live in complex 3-dimensional tissues, interacting with each other, sending signals, influencing each other’s growth. They have neighbourhoods with infrastructure and, as we all know, your neighbourhood can decide who you are. Life is 3-dimensional.
3D is the real deal. Not only on the screen but also in cell culture dishes.
No wonder labs are abandoning petri dishes and turning to 3D cell culture. So how is it done and is it worth the effort? Should I go 3D? Where is it all leading?
There are three main forces driving the current shift from 2D to 3D. The first is realism. Cells just don’t act naturally when you take them out of the environment. And let’s face it, dropping a dimension is about as unnatural as you can get. Losing a dimension changes everything – Google Earth looks better than Google Maps.
The second driver for change is partly ethical: 3D culture is seen as a bridge between in vitro and whole-animal approaches, to some extent replacing the latter. And thirdly, there is increasing confidence that growing cells in 3D is a route to creating replacement tissues in therapy.
If you are a 2D die-hard, just stop for a moment and think about the mechanics. Cells growing on a surface change their shape as they grow, flattening out, perhaps eventually making contact with each other until they almost cover the surface. At this stage they look like fried eggs – half of their membrane touches the surface, only half in contact with the bathing medium.
That restricted access to medium is a problem in itself. But there’s worse. A flat plane of cells means only a tiny fraction of their membranes are in touch with each other. Subsequently, one vital route of communication – direct cell-to-cell contact – is virtually wiped out. A cold handshake in place of a warm hug. The cells are almost in solitary confinement; at least as far as direct contact goes. And it is not as though the other routes, such as chemical signalling, are okay either – see below.
As cells develop, cell contacts can influence several aspects of the cell’s phenotype. Cell contacts play a big part in what shape the cells will adopt and how they behave. Think of epithelial cells, and how their physiology depends on their polarity. This is often entirely dependent upon having contact with the appropriate cells or basement membrane. And then again there are neurones – many of them simply refuse to send out processes without contact with other cells, and will almost never form synapses unless you culture them with glial cells. Neurones without synapses? Just how realistic is that?
Whenever cells touch each other, they affect each other through cell surface receptors and these receptors have a strong regulatory effect on specific patterns of gene expression. To put it bluntly, isolate a cell physically from other cells and your cell turns, like Dr. Hyde, into something else. As the poet once warned, “No cell is an island”, or something like that. This is bad news. After all, the main reason for doing cell culture in the first place is because you want to use your cultured cells to find things out about real tissues.
This was brought home by a key experiment in 1992 by Bissell and colleagues (Petersen et al., PNAS, 89, 9064-68). This group was trying to grow human breast epithelial cells in the usual 2D way but, instead of turning out as normal breast epithelial cells, they insisted on growing as cancer cells. But when they turned to culturing the cells with reconstituted basement membrane, they found they could get normal cells in culture. In other words, until they were given a 3D environment, the cells just wouldn’t act as themselves.
Keeping in contact isn’t the whole story. Much of the oddness of 2D cell culture is the very thing that initially attracted us: it is so, well, samey. All the cells get the same medium, all get the same rate of flow of medium, all are subject to exactly the same concentrations of signals, oxygen, CO2 and so forth. All the cells are neatly laid out in a uniform flat array. Very convenient: we can apply drugs and know all the cells see them and it was great for imaging, especially with early microscope technology.
Stepping up to 3D means you can mimic many other aspects of cells’ natural environment and it provides cues, which are essential if the cell is to behave like it would “at home”. In real life, it is the very inhomogeneity of the environment that makes cells work together. Cells get in the way of diffusing materials and this sets up concentration gradients that are vital in intercellular signalling. And in real life, unlike on a flat surface, cells experience limitations to motility – social mobility is as much a part of a cell’s life as it is for us.
In one sense, 3D culture is not entirely new. Many researchers found themselves up against a dead end in their work because of cells that simply didn’t behave properly when grown in culture – their shape was wrong, or they lost some key aspect of their phenotype, or perhaps wouldn’t even grow in the first place. So, they naturally wondered if there was some key aspect of their environment that was missing. As a result, it was commonplace to cover the growing surface with collagen (or, if you could get away with it, a cheaper mixture of less pure forms of it, called gelatin).
It may surprise you to know that the first step to real 3D culture began as early as the 1950s. Remember filter-well inserts? Cells grown on filters that could be suspended in mid-media, as it were. This meant you could have different media on either side, introducing polarity into cell culture for the first time. It was soon spread around that epithelial cells grew much better like this, and filter-well inserts became a standard commercial product.
So how is it done? You could create the simplest form of 3D culture without any new equipment. Enter the hanging drop. As the name suggests, this is a matter of suspending cells in a droplet of medium hanging from a surface. The curvature of the droplet and the absence of an adhesive surface, encourages the cells to develop as a clump. Many cells will naturally adhere to one another in these circumstances. It is cheap and technically easy but has a number of disadvantages. The cells on the outside of the clump will have free access to the medium and therefore to nutrients, and will be able efficiently to exchange oxygen and carbon dioxide. The cells in the middle of the clump, however, fare worse and can often form a core of dead and dying cells. After all, cells in active tissue are rarely more than 100 micrometres from a rich oxygen supply.
Or you can try a hydrogel. Hydrogels are matrices of proteins, in which gaps between the protein components form pores where the cells grow. It is important to get the pore size right, of course. Hydrogels range from completely natural in origin to completely synthetic, most being a mixture of both. The best known is the commercial Matrigel, which is a reconstituted basement membrane collected from the Engelbreth-Holm-Swarm mouse tumour. But take a look around and you’ll find a huge array of alternatives, including old favourites like collagen and hyaluronic acid, and more exotic ones, like those based on silk proteins.
Of course, natural hydrogels are essentially just extracts of the cell’s natural environment, so they contain many of the growth promoting elements of the source material. But herein lies an element of danger, or perhaps two... Firstly, they are complex and ill-defined, and so can vary in composition, making it difficult, or impossible, to keep consistency between batches. Secondly, you are stuck with what you start with – you don’t have the option of fine tuning the composition to the specific needs of your culture. Okay, three danger elements: if you consider contamination risks, particularly from viruses.
So are synthetic hydrogels any better? At least they mean you won’t have problems with consistency or tuning – you can have things all your way by adjusting details of the manufacture. And you can incorporate extra elements, such as extracellular matrix proteins (laminins, proteoglycans), poly(lactic acid) or poly(caprolactone) units into the backbone. You can make (or buy) gels with different viscosity or other physical properties. Many synthetic hydrogels are designed to degrade slowly, so that cells can lay down their own extracellular matrix, migrate, or change their morphology. Indeed, controlling how stability is maintained is not trivial and commercial suppliers are actively addressing this issue in developing new gels.
A key feature of all hydrogels is that they contain heavily cross-linked proteins, which, with their negative charge and an ability to coordinate ions, allows for a high water content. This means they are highly permissive to diffusion, being about 99% water. Because of these advantages, the development of synthetic hydrogels is proceeding apace. More recent ones consist of self-assembling proteins that automatically (in the presence of key agents, such as specific salts) form gels. The regular structure of the peptides, including regularly-spaced side chains, means the size of the pores can be tightly controlled.
Cells cultured in hydrogels form isolated clumps. This gives three dimensions to the culture, so in this respect, life in the clumps is fairly normal, even if the clumps never talk to each other. It is easy to understand, then, why this approach has proved a favourite in research into tumours. But if you are serious about 3D culture and, in any case, with the need to get beyond the size limitations of gels and drops, you need a scaffold.
And when it comes to the type scaffold, the sky is the limit. But be warned: successful 3D culture means choosing the right one. And there are many, many issues to consider. What about the biological compatibility of the material? You don’t want anything in the “bulk chemistry” – what the scaffolding material is made of – to harm the cells. And it must be stable, so that there is no degradation over the life of the culture. Then you have to think about the “surface chemistry”: the properties in the surface of the material, which, after all, is what the cells will be in contact with. Where the bulk chemistry is a protein, this will usually be determined largely by the charge and polarity of the surface material.
And don’t forget what you are going to use the culture for. Are you going to apply any ligands that might meet with access issues? What about delivery of stains or dyes for imaging? Will you need to be able to control the rates of diffusion? Culturing on scaffolds makes a whole class of new stimuli possible, such as stress, shearing, pressure and tension. Will the scaffold bear these stimuli without breaking?
Choosing the right scaffold material and architecture is a prerequisite for successful 3D cell culture: osteoblast growing on scaffold made of calcium oxide and silicon dioxide.
Faced with this bewildering set of needs, the past decade has seen rapid strides in methods for making better scaffolds. Electrospun fibre scaffolds (think of really, really small candyfloss machines) offer good control of pore size, by altering parameters of the spinning, such as the flow rate, collecting distance and the properties of the polymer solution. Matrix molecules can be incorporated and this has been shown to improve cell growth (indeed, even spinning collagen is better than growing cells on it). The size of the fibres, which also affects cell growth, can also easily be controlled. On top of that, you also have fine control over the “nanotopology” – the way in which the fibres lie in relation to each other. This can revolutionise neuronal culture because many neurones will happily grow axons along a spun fibre scaffold, if the fibres are arranged in parallel.
So it won’t surprise you to hear that advances in 3D cell culture go hand-in-hand with the rapidly developing field of nanotechnology. What we are already calling “traditional” methods, such as solvent casting, particle leaching, salt fusion and gas foaming, are already being replaced with revolutionary new techniques. “Rapid Prototype Processes” refers to a new class of automatic manufacture, in which you can design any complex pore geometry.
For example, Engelmayer et al., used lasers to cut pores in PGS (poly[glycerol sebacate]) membranes to produce tailor-made blocks that could then be assembled layer-by-layer into a scaffold (Nature Materials 2008, 7, 1003-10). Culturing heart cells in this scaffold resulted in a tissue much more like heart cells were properly aligned and they could even induce contractions.
But before you ditch all your petri dishes, you should be aware the field is not without major challenges. The biggest hurdle remains that bane of all culturing – difficulties of automation. 3D culture is just as much a black art as 2D culture and, whether you are a small lab or a big drug developer, you will have to spend time looking for the right combination of factors to get things working.
It is just that the problems move up to 3D, too. The use of cultured tissues in therapy is severely sensitive to legislation. Controlling for the effects of diffusion and access is difficult. But the exploding rate of 3D publications and the expansion of 3D culture materials offered by major suppliers are revealing, and already microplates with built-in scaffolds are being developed for use in drug discovery.
Suddenly, growing cells on a flat sheet of plastic is beginning to look a bit silly.
Last Changed: 07.02.2013