Cell Cultures and Laser Microdissection

Many of the discoveries that are now being made in cell division and differentiation, the relationships between single cells and cell organelles, treatments of cells with pharmaceutic substances, etc. would not be possible without live cell cultures. Allowing morphological and biochemical observations of single cells under different experimental conditions, they provide a unique source of information.  The challenge with cell cultures is to visualize them by using suitable contrast techniques without killing off the cells.


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Applications of laser microdissection for live cell cultures

Laser microdissection offers a wide range of manipulation and selection options for cell cultures, isolating not only single living cells but also whole cultures from homogeneous primary cultures for further processing or cultivation.

Single cells or cell aggregates can be easily microdissected using various Live Cell Cutting (LCC) modules which dissect selected areas or distinct clones from cell cultures for further cultivation or molecular-biological analysis such as PCR. To guarantee the required sterility and undamaged condition of the cells, they are often already cultured in special petri dishes or specimen slides with special wells for cell cultures (www.ibidi.com). These are placed on the specimen stage in specially designed holders and then observed, manipulated and dissected with the laser microdissection technique. A distinction is made between conventional petri dishes and petri dishes with PEN membranes. The latter are particularly suitable for the live cell sector using LCC. After laser microdissection, the dissected cells fall, for example, into 8-well strips which can be put on 96-well racks in the incubator and thus recultivated.

The possible uses and combinations of the various specimen and collector holders in laser microdissection are described in the following sections.

Fig. 1: Human forescin fibroblasts infected with human cytomegalo virus, HCMV-GFP fusion protein (Courtesy of: Margarete Digel and Dr. Christian Sinzger, Institute of Medical Virology, UKT University of Tübingen, Germany).

Possible combinations of specimen and collector holders for laser microdissection

The following specimen and collector devices can all be used and combined in LCC mode: petri dishes with PEN membranes, stackable membrane rings and Ibidi slides. These three specimen carrier can be used for laser microdissection and selection, and are also suitable as collection devices. 8-well strips and petri dishes (without PEN membrane) can also be used as collection devices, but cannot be used for laser microdissection. However, live cell cultures and single living cells in petri dishes without PEN membrane can be manipulated with the laser and watched over longer periods of time using the time-lapse movie function.

In particular, all specimen devices suitable for cell culture are combinable with all customary collection devices (0.5 ml tubes, 0.2 ml tubes, 8-well strips, Lab on a Chip (LOC), Ibidi slides and petri dishes).

Petri dishes with PEN membranes
Cells cultivated in petri dishes with PEN membranes can be dissected in LCC mode and transferred to other petri dishes (with or without PEN membrane). Equally, the dissectates from the petri dishes can be directly collected in so-called Ibidi slides and 8-well strips for further cultivation or in 0.5 ml tubes, 0.2 ml tubes, 8-well stripe caps, or Lab on a Chip (LOC) for molecular biological downstream analysis.

Stackable membrane rings
Stackable membrane rings offer the same possibilities as petri dishes with PEN membrane. In addition, they can be stacked underneath each other for laser microdissection to capture isolated material directly in the downer ring from the one above.

Ibidi slides
Ibidi slides are specimen slides with special wells and, like the above, are combinable with the entire range of specimen holders suitable for the LCC mode as well as with all customary collection devices for downstream analysis (0.5 ml tubes, 0.2 ml tubes, 8-well strips and Lab on a Chip (LOC)). In addition, an Ibidi stack is an elegant, contamination-free selection method: If cells from such a stack of specimen holders are dissected, the dissectates fall naturally into the next slide below, which is held in a special stack holder, for recultivation.

Recultivation after laser microdissection

After microdissection, the cells pass to one of the collection devices and can be recultivated in a fresh nutrient medium.  Stackable membrane rings do not require a special collection holder. These and Ibidi slides can simply be stacked upon each other allowing a contamination free microdissection process. However, in combination with a suitable cover (ibidi slides) or optional climate chamber for the laser microdissection system, a contamination-free selection process is guaranteed. Another advantage of stackable membrane rings and Ibidi slides is that when a cell is dissected, no separate collection holder is needed.

Also, all customary collection devices such as Eppendorf caps or Lab on a chip (LOC) can be used for molecular-biological analysis and PCR, for example, without any further preparation.

Practical use of live cells in laser microdissection (LMD)

One of the aims in basic research is to find ideas for a new cancer drug. This example is described in the following.

In the experiment, tumor cells are cultivated in the form of a primary culture and split before reaching confluence. Some of the cells are recultivated in order to obtain enough cell material for further experiments, while the others are applied to a petri dish with PEN membrane.

For the further procedure, the nutrient medium in the petri dish first has to be removed until only a thin film remains. As a petri dish with PEN membrane is used, single cells in the existing culture can easily be selected by LMD in LCC mode and transferred to 8-well strips. There is now a single tumor cell in each well  

The procedure can be repeated until each well of the collection device contains an individual cell again, which in this case forms the basis of a new culture after dissection. The cells of each culture of a well should therefore contain the same DNA, as they stem from one and the same isolated original cell, and the cells can be lysed and forwarded for PCR analysis for mutation detection. Results are thus always traceable to the dissectate of one single cell.

Fig. 4: Laser microdissection and living cell culture (LCC). Experimental set up for the benefit of laser microdissection (LMD) for cultivated cells. A) shows the routine procedure of cultivating and splitting cells (in this example a Tumor culture with heterogeneous cells): after confluent growth cells are splittet and re-cultivated into usual petri dishes for further cultivation and into petri dishes with PEN membrane for further processing with LMD. B) line outs an example of specific cell extraction into 8 stripe wells, enabling investigations of cell growth under different conditions or of different cells each well. C) Recultivated cells are selectively selected with LMD for cloning: Single tumor cells (turquoise) are dissected into wells of an Ibidi slide for further cultivation while unaffected cells (grey) remain in the petri dish. Ibidi slides can be used as a stack ("sandwich") for laser microdissection to isolate cloned cells into several wells of different Ibidi slides for multi parallel analysis like treatment with different toxins.

Besides this, further cells of the primary culture can be regularly transferred to a petri dish with PEN membrane on splitting. LMD can be used to dissect new single cells from these tumor cells in the petri dish for recultivation in another petri dish with PEN membrane. Again, single cells of the same original cell can be isolated in LCC mode, analyzed by PCR as described above and transferred in parallel to Ibidi slides. Toxins can now be added to these in order to analyze their influence on tumor cell growth in comparison to cell growth of healthy cells (Figure 2).

With regard to the development of a new cancer drug, different toxins could be applied to cells of the same origin with identical DNA in order to examine the reaction of the cells to particular toxins. It is also possible to observe the genome of the cell (and the gene and protein expression, if desired) via molecular-biological techniques.

A further possibility is to treat different single cells with different DNA of the same tumor with the same toxin. The reaction of the various single cells to the applied toxin can then be examined and compared.

Instead of a conventional counting chamber or manual selection of many cells, an Auto Vision Control (AVC) program can be used, which identifies and selects cells for laser microdissection on an automatic basis. This implies a large throughput of samples and therefore enormous time savings. The AVC module is a particularly viable alternative to the manual acquisition of individual cells in protein research. The AVC program is also able to capture and select fluorescing cells. For the successful transfection of foreign DNA into eukaryotes, a fluorescence marker can be coupled to the introduced vector as a control. For example, GFP is infiltrated via a vector; the labeled cells containing GFP proteins after successful infiltration and further cultivation fluoresce and can be easily counted and dissected.

Fig. 5: Lung epithelial cell culture.

Use of cell cultures in research

Cell cultures are being increasingly used in medical research as well as in the pharmaceutics and cosmetics industries and in biotechnology and environmental analysis. A major application is in basic research for the study of metabolism, division and other cell processes. They are also useful as test systems, e.g. for examining the efficacy of substances for signal transduction and cytotoxicities.

Simple proteins can be easily produced in bacteria, whereas more complex proteins can only be produced with a cell culture to enable the right protein glycosylation and complicated linking via disulfide bridges to take place.

Live cells also play a significant role in the manufacture of various biochemical products such as monoclonal antibodies for research and therapeutic application in medicine. A few of the many possible uses of monoclonal antibodies are therapies for acute and chronic leukemia, radio-immune therapies of non-Hodgkin lymphoma or the treatment of rheumatoid arthritis. Certain vaccines (for instance against influenza) can also be produced from cell cultures. And erythropoietin, applied as a hemopoietic treatment in connection with breast cancer, for example, is also obtained from cell cultures.

Nearly all cell types can be generated from stem cell cultures. One familiar practice is the use of hemopoietic stem cells in leukemia research and therapy. Adult stem cells can also be used for tissue engineering.

In diagnostics, potentially pathogenic microorganisms are applied to cultures and analyzed after proliferation has taken place.

In plant research, complete plants can be generated from undifferentiated meristematic cells. For example, Arabidopsis could be mass-produced in vitro to perform tests on salt stress, water tolerance, etc.

So-called bioreactors are found in industry in particular for working with live cells – a typical application is the production of modern insulins. Mesenchymal stem cells can be used for the regeneration of various types of tissue, as they can be differentiated in various somatic cells such as cartilage, bone or muscle. To obtain differentiation in the particular cell type, tissue-specific bioreactors are used, e.g. to develop a vascular skin equivalent for use in in-vitro examination of malign tumors. Similar experiments with standardized human cell material are to serve as an alternative to animal experiments in future or to replace them altogether.

All the above examples, however, only show a small selection of a huge range of possibilities and opportunities in live cell research and development.

What are cell cultures?

A cell culture is the cultivation of animal or plant cells outside the organism (in vitro) under controlled conditions in a nutrient medium. A distinction is made between primary and immortalized cell cultures. A primary culture is a non-immortalized cell culture taken directly from a tissue sample. An immortalized cell culture, on the other hand, is characterized by its unlimited capacity for division. This can take place either by random mutations, e.g. in tumor cells, or by deliberate genetic changes such as the introduction of a dedicated plasmid vector. Such cells can display characteristic features of an in vitro neoplastic or malign transformation.

Basically, single cells can be obtained from all kinds of tissue (e.g. liver, skin, kidney, brain cells, etc.). This is done by using proteases such as trypsin which breaks down the proteins that keep the cells together. The cells isolated in this way are then cultivated and multiplied in petri dishes. Multiple division can be selectively stimulated in some cell types by adding growth factors.


Unlike bacteria, most animal cells/primary cultures are not able to survive in suspensions and need a suitable adhesion surface for their growth. It is therefore possible to differentiate between adherent cell cultures (those that grow on the surface such as fibroblasts, epithelial, endothelial and cartilage cells) from non-adherent suspension cultures, in which the cells multiply as they swim freely in the nutrient medium, such as lymphocytes.

As cells divide in a suitable culture medium, a so-called cell lawn is formed. When adherent cells attain the greatest possible density in their arrangement, we speak of confluence. This triggers a contact inhibition which stops the cell division and the culture becomes stationary. Vertebrate cells usually only divide 30 to 50 times before apoptosis. Cell immortalization is induced by sometimes spontaneous but usually experimentally induced measures such as the introduction of a special plasmid as vector. A primary culture thus becomes a permanent cell line which is subject to an unlimited division rate but has a number of characteristics of its original cells.

Transformed malign cells usually lack the contact inhibition phenomenon, presenting themselves as immortalized in culture and surviving in suspensions. As they have the characteristics of tumor cells, tumor formation can be induced by injecting them into an organism/experimental animal.


Nutrient media must be such as to ensure the growth, proliferation and differentiation of the cells that are being multiplied as well as the execution of the typical functions of the cell. The optimal composition depends on the particular cell type. Chemically defined media are generally preferable; typical basal media contain, besides amino acids, vitamins, inorganic salts and buffer mixtures, additional growth and differentiation factors that can be replaced by fetal calf serum.

Cultivated cells only survive in the presence of suitable media. The pH value must be monitored, generally by a color indicator in the nutrient medium. Cell proliferation normally takes place at a temperature of 37°C with an atmosphere of 5% CO2 as buffer for the medium, in special incubators. After they have multiplied, cells can be removed from the surface of their petri dish either chemically (e.g. trypsin) or mechanically (e.g. cell slide) and sub-cultivated in a suitable dilution. An adherent cell culture should be passaged i.e. transferred to a fresh nutrient medium, before confluence occurs. In this way, primary cultures can be kept alive for relatively long periods.

Methods for visualizing cell cultures

Observation of living material is still a prerogative of light microscopy. Modern high-tech instruments such as scanning electron microscopes prove unsuitable despite their high resolution.

However, live cells are by nature transparent in the visual spectrum and therefore exceedingly poor in contrast. Under a conventional light microscope, an almost “transparent”, hardly recognizable image is produced in the so-called brightfield mode.  Although there are many specimen staining products for obtaining the desired contrast, cell cultures are usually killed off by preparation and staining processes.

Live cells can be imaged with the phase contrast technique, although crosstalk and distortions at thicker parts of a specimen cause problems here. Differential interference contrast methods (DIC) are another alternative for imaging cell cultures and live cells. The Integrated Modulation Contrast (IMC) technique has also proved useful for examining unstained, low-contrast samples. Another established method of visualizing live cells is to label them with fluorescent proteins. These can be genetically introduced into bacteria and into animal or plant cells. The method can be used for labeling single live cells, cell aggregates, organs or whole animals. Protein biosynthesis as well as transport, secretion and degradation processes can be directly observed and analyzed in organisms using fluorescence. Fluorescing proteins exist in many different forms with different fluorescence spectra, e.g.

  • GFP (Green fluorescent protein)
  • BFP (Blue fluorescent protein),
  • CFP (Cyan fluorescent protein),
  • YFP (Yellow fluorescent protein).

Luciferins and fluorescing coral proteins are also being increasingly used in microscopy.  GFP is routinely applied to label proteins in live cells and examine their behavior in situ.

Digression: Fluorescence/GFP

Some substances emit light when exposed to short-wave radiation. In fluorescence technology, this is utilized by labeling non-autofluorescent structures of interest with appropriate fluorescing proteins such as GFP. GFP is the modified protein of a bioluminescent type of jellyfish (Aequorea victoria). When excited with blue or ultraviolet light, it emits an intensive light-green color.

When introduced into cells by genetic engineering techniques, fluorescing proteins give insights into the distribution, anatomy and development of certain cell forms. As they have no toxic effect, they are extremely suitable for exploring live cells. The cells are neither damaged nor destroyed, so conditions are ideal for experimenting on live organisms in vivo, in situ and in real time.

How does the genetic information get into the cell?

The answer to this question is that special transport plasmids are introduced to the target organisms as vectors. The fluorescent protein is normally coupled to these.

The vectors work in different ways:

Cloning and expression vectors stay in the target organisms even after fulfilling their transport task. The suicide vectors that can be introduced into many hosts are not able to establish themselves in these. Because of their transposomes (so-called jumping genes) the suicide vectors manage to penetrate into chromosomes or plasmids of the target organisms, taking the coupled GFP with them. The rest of the suicide vector is lost.

Cell cycle arrest vs. FUCCI technology

Another technology that could also be combined with the use of laser microdissection in future is FUCCI (Fluorescent Ubiquitination-based Cell Cycle Indicator method).

In this method, cell lines are specifically treated to change their fluorescence according their cell cycle state. Proteomic studies could be carried out in situ, in vivo and in real time during the cell cycle to capture development stages of a distinct cell with regard to its protein expression and to dissect desired cells or areas using laser microdissection. Furthermore, it is also possible to select individual cells of the same cell cycle phase quickly and directly using AVC software, for example.  

Cell cycle arrest

So-called checkpoints are particularly important for “quality-checking” the cell cycle during the phase transitions. It is possible to arrest the cell cycle  at the G1-/S-phase (metaphase). Up to now, it proved difficult to determine the exact time and place of the  phase transitions – it could only be done by a chemically induced cell cycle arrest process, for example. Capturing live cells of a certain stage is problematic as cells have different division rates and are therefore at different phases of the cell cycle.

If a toxin is applied to the cells before the S phase so that division stagnates, the cell cycle can be synchronized within 2 days. All cells are then in the same phase. This type of cell cycle arrest is used for isolating chromosomes of leucocytes, for instance. Leucocytes of human blood are cultivated over a period of 72-96 hours and then treated for one hour with colcemide to halt the cell cycle in the metaphase.

However, the synchronized cell cycle becomes imbalanced again after a while due to the different division times.

FUCCI technology

FUCCI technology can be specifically used to visualize the transition from G1 phase to S phase in live cells. The problem of long waiting periods as encountered with the chemical induced cell cycle arrest technique is circumvented by introducing fluorescent proteins. The different stage of development of the cells can then be identified by their different colors. Fluorescence is triggered by introducing the proteins mAG-geminin and mKO2-Cdt1, for example. The individual phases are identified by striking color contrasts. Cell nuclei show up red in the G1 phase, and fluoresce green in the S-, G2-, and M phase. The transitions from G1 to S phase are identified by yellow fluorescence. The combination of fluorescence mode with laser microdissection is an enormous time saving, as the yellow fluorescing cells, for instance, can be captured via the AVC module and directly recorded and dissected in fluorescence mode.