A Clearer View of 3-D Cell Culture: Optimizing Fluorescence Imaging and Analysis

The study of cells in two-dimensional (2-D) culture systems, usually in a well plate or in a flask, has contributed to the understanding of many biological processes. Monolayer culture has been the protocol of choice, from investigating cell and molecular biology in adherent cultures to developing predictive models of therapeutic drug action.

The 2-D monolayer method is limited in that cells do not exist in an isolated 2-D environment in vivo. Instead, they are surrounded by cells of different types, where they are exposed to signaling chemicals and their respective gradients. They are also typically embedded within the extracellular matrix (ECM), which is capable of inducing cellular changes.

Three-dimensional (3-D) culture systems were developed to address these issues. Culturing cells in 3-D is not new, but investigation highlights a range of advantages relative to 2-D culture. For example, cell fate and differentiation can be heavily influenced by the way in which cells are cultured. This has led to the idea that 2-D cultures can actually drive abnormal cell function or de-differentiation, confounding results and calling into question the physiological relevance of the data generated.¹

The ECM plays an important role in regulating the distribution of nutrients, gases and various chemicals (e.g., morphogens, cytokines and hormones) as well as helping to define cell shape and organization. Cells cultured in 2-D are not completely surrounded by this matrix. As a result, they tend to have altered morphologies. Monolayer cultures force cells to assume apical–basal polarity, an unnatural state for some cell types—mesenchymal cells, for example, which would normally only polarize from front to rear during migration.²

Two-dimensional culture models have been particularly restrictive in the field of cancer research, where the culture of tumor spheroids (tumoroids) has been pivotal in allowing researchers to generate biologically relevant data, providing insight into tumor structure,^3,4 therapeutic response prediction, ⁵ the role of hypoxia⁶ and even drug penetration.⁷

The need for visualization

While 3-D cultures more accurately mimic the in vivo environment, it can be very difficult to image cells in 3-D. The tight structure of spheroids, the lack of transparency of many plastic materials or the dense fiber network of some hydrogels can interfere with detection of the fluorescent signal. Since fluorescence imaging is widely used to analyze cells in culture, this can be a significant obstacle.

Getting the most out of 3-D immunofluorescence

Researchers are now turning to advanced systems to conduct immunofluorescence image analysis of 3-D cultured cells without having to use an algorithm-based correction method driven by deconvolution software. These systems typically include all of the reagents and consumables required to produce 3-D cultures that can be effectively imaged under a standard widefield microscope.

Two different cell types were used to assess the quality of imaging achieved with the RAFT 3D Cell Culture System (Lonza, Walkersville, Md.). Primary human neonatal dermal fibroblasts (HDFs) and cells from the breast cancer cell line MCF7 were cultured under similar conditions. HDFs were seeded at 5000 or 50,000 cells per well, while MFC7 cells were seeded at 10,000 cells per well, both in 96-well pates. The cells were then fixed with 3.7% formaldehyde solution after three days (HDFs at a starting density of 50,000 cells/well), 11 days (HDFs at a starting density of 5000 cells/well) or 23 days (MCF7 cells) of culture at 37 °C, with 5% CO₂.

Subsequently, the fixation was replaced with 100 μL of quenching solution (1 mM tris-HCl and 20 mM glycine in phosphate-buffered saline [PBS]) to quench the formaldehyde cross-linking, and the plate was incubated at room temperature for 10 minutes. After washing the RAFT cultures three times for 15 minutes with 100 μL PBS, the PBS was replaced with either 100 μL of 0.1% Triton X-100 (Sigma-Aldrich, Saint Louis, Mo.) solution (HDFs) or 1.0% Triton X-100 solution (MCF7) in PBS to permeabilize the cells, and the plate was incubated at room temperature for 4 minutes.

The cells were first stained for tubulin using YOL1/34 rat anti-tubulin primary antibody (Abcam, Cambridge, U.K.) diluted in 1% (w/v) bovine serum albumin in PBS (with 0.2% Triton X-100 for the MCF7 cultures) and visualized with Cy3-AffiniPure Goat Anti-Rat IgG secondary antibodies (Stratech Scientific, Suffolk, U.K.; in the U.S., Jackson ImmunoResearch Laboratories, West Grove, Penn.) in the respective blocking buffer. Cells were also stained with Alexa Fluor 488 phalloidin (Life Technologies, Grand Island, N.Y.) and 4’,6-diamidino-2-phenylindole (DAPI) (Life Technologies) in order to view the actin filaments and nuclei, respectively. Primary antibodies were incubated at 4 °C overnight, and secondary antibodies were incubated at room temperature for 2.5 hours. The cells were then imaged on a widefield IX71 inverted microscope (Olympus, Center Valley, Penn.) with 10× and 40× phase contrast objectives. Fluorescence from the antitubulin antibody and the phalloidin was strong enough that imaging at an exposure of only 100–200 msec was required.

This experiment yielded high-quality immunofluorescent images of HDFs displaying typical elongation and microtubule cytoskeletal structures (Figure 1), and MCF7 cells forming the expected rounded morphologies (Figure 2). To compare the data to the standard approach using deconvolution, the researchers used parallel spectral deconvolution on fixed HDFs (Figure 3). Microstructures within both the HDF and MCF7 cultures could be seen without the need to apply deconvolution (Figures 1 and 2).

Figure 1 – HDFs fixed and stained for tubulin and actin after being cultured in the RAFT 3D Cell Culture System for three days. A series of z-planes taken at 0.5-μm intervals was imaged on a widefield microscope after staining the cells for tubulin (red), actin (green) and nuclei (blue). The z-stack from each channel was projected onto one plane using the maximum z-projection function in ImageJ open-source software, and the merge of all channels is shown in the large bottom panel. Across the top panel, one frame of the z-stack is shown, with each individual channel represented in gray scale separately to better display the detail of each staining.

Figure 2 – MCF7 tumoroids fixed and stained for tubulin and actin after being cultured in the RAFT 3D Cell Culture System for 23 days. One z-plane was imaged on a widefield microscope after staining the cells for tubulin (red), actin (green) and nuclei (blue). In the larger images around the exterior, each channel (labeled directly) is shown individually in gray scale, and then merged in the bottom right-hand corner in color. In the center panels, a dividing cell indicated by a white box on the larger images is shown for each channel and the merge at a higher magnification.

Figure 3 – HDF fixed and stained for tubulin after being cultured in the RAFT 3D Cell Culture System for 11 days. A single z-plane was imaged on a widefield microscope after staining cells for tubulin (red) and the nuclei (blue). The image from each channel was deconvolved using the 3-D spectral deconvolution software from ImageJ (Generalized Tikhonov) and the merge of the two channels is shown.

The data also indicates that the RAFT Cultures allow the permeation of antibodies (typical molecular weight of 150 kDa). Therefore, standard 2-D immunofluorescence protocols can be used with RAFT 3-D Cell Culture. For cellular aggregate structures such as the tumoroids formed from the MCF7 cells, a higher concentration of the permeabilization reagent may be required—1.0% Triton X-100 compared to 0.1% Triton X-100 in PBS. In addition, use of a 1% bovine serum albumin (BSA)-containing blocking solution for antibody dilution is recommended to decrease potential background noise from nonspecific antibody binding to the collagen matrix. Interestingly, no cross-reactivity of the goat anti-rat IgG secondary antibody with the RAFT Culture Matrix was observed. This indicates that the collagen does not contain any detectable amounts of rat IgG and confirms the high purity and quality of the rat-tail collagen type I provided in the RAFT reagent kit.

Conclusion

A 3-D culture system more closely mimics the in vivo environment than traditional 2-D methods, allowing cells to form complex cell–cell and cell–matrix interactions. Advances in 3-D systems are facilitating the creation of more physiologically relevant models that will ultimately lead to an improved understanding of cell biology.

References

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5. Vinci, M.; Gowan, S. et al. Advances in establishment and analysis of three-dimensional tumor spheroid-based functional assays for target validation and drug evaluation. BMC Biol. 2012, 10(1), 29; doi:10.1186/1741- 7007-10-29.
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Dr. Cecile Villemant is Development Scientist, and Dr. Grant Cameron is RAFT Development Director, TAP Biosystems, York Way, Royston, U.K.; tel.: +44 1763 227200, fax: +44 1763 227201; e-mail: [email protected]; www.tapbiosystems.com. Dr. Jenny Schroeder is Senior Scientist, Lonza, Cologne, Germany.