C ELL CULTURES TECHNIQUES

All aseptic cell cultures procedures were carried out in a dedicated cell culture laboratory within a laminar flow cabinet (Aura 2000, Bioair Instruments, Pavia, Italy). Glassware was sterilised in an oven at 180 °C for 3 hr prior to use and cell culture media was warmed to 37 °C before use. Cell culture materials were purchased from Sigma-Aldrich (Poole, UK) and all cell culture disposal plastics were from Fisher Scientific (Loughborough, UK) unless otherwise indicated.

3.4.1 Endothelial cells

Human Umbilical Vein endothelial cells (HUVECs) were chosen as a cell model to investigate the interaction between cells and the polystyrene surfaces modified by UV/Ozone treatment. HUVECs were purchased from European Collection of Cell Cultures (ECACC) and propagated from a cryopreserved immortalised cell culture originally isolated from normal human umbilical vein. HUVECs were used in this study between passages number 60-70. Cells were normally cultured in Dulbecco’s modified eagle’s medium (DMEM, D5796, 4.5µg/mL glucose, L-glutamine and sodium bicarbonate) containing 10%

foetal calf serum (FCS) and 1 unit/ml penicillin, 1 µg/mL streptomycin and placed in a constant atmosphere incubator (Galaxy S, Wolf Laboratories, York, UK) at 37°C with 5% CO2 and 95% air known as normoxia. Similarly, the combination of 5% CO2, 5% O2 and 90% air (not including O2) is for HUVECs cultured under hypoxia condition, nitrogen was inflated into the incubator to reduce the oxygen concentration to create the hypoxic atmosphere.

3.4.2 Cell resuscitation

HUVECs were purchased as a cryopreserved vial which contained 3.75 x 10⁵ cells/mL. The vial was thawed by dipping one third into a 37°C water bath and the cell suspension from the vial was transferred to a sterlise flask (75cm²) containing 10ml of cell culture medium. After incubation for 24 hr at 37°C with 5% CO2, the cell culture media was replaced to remove any cell debris, unattached cells and any traces of dimethysulfoxide (DMSO). Cell culture media was replaced every two days until cells reached 80%

confluency. Cells were then split prior to setting up specific experiments or for long term storage.

60

3.4.3 Cell passage

Cell passage or cell splitting is the process of sub culture of cells in order to allow cells to increase in number in specific conditions. To sub culture cells, growth medium (DMEM with 1% FCS, 1 unit/mL penicillin, 1 µg/mL streptomycin) was discarded and the flask was washed with phosphate buffered saline (PBS) three times. The flask was then incubated for 10 min in the final wash to remove any medium containing serum. Trypsin was used to remove the adherent HUVECs from the culture surface; 5ml of 0.25% (v/v) trypsin solution was added to the flask and incubated for a further 10 min to detach the cells.

The flask was checked using the optical microscope to observe cell detachment. Following detachment of the majority of cells from the surface, an equal volume of growth medium containing FCS was added to the flask to neutralize the enzyme activity of the trypsin which may damage the cells if left in contact for longer times. The cell solution in the flask was then transferred to a sterile centrifuge tube and spun in a centrifuge (Biofuge PrimoR, Heraeus, Germany) under 220g force for 8 min. After centrifugation, the supernatant was carefully discarded into a waste beaker and 1 mL of fresh growth medium was added to the cell pellet. Cell counting was then performed as described in Section 3.4.4 and, for routine subculture, a cell suspension containing 5 x 10³ cells/cm² was transferred to a new flask with 10 mL of fresh growth medium and left in the incubator to allow the cells to attach and proliferate. The flask was checked daily under the optical microscope to monitor progress and potential contamination.

3.4.4 Cell counting

The counting of cells was carried out using an improved Neubauer cell haemocytometer (Weber Scientific International, West Sussex, UK) under an optical microscope. Briefly, trypsinised cells were collected by centrifugation, and washed once in PBS. Cells were re-suspended in 1 ml of medium and a small volume was incubated with an equal volume of 0.4% (v/v) trypan blue solution and transferred to the haemocytometer. Trypan blue is a vital staining method to selectively stain dead cells. Live cells with intact cell membranes were not coloured since the dye is unable to diffuse into the cell via the cell membrane. However when the cell membrane is damaged, typical of dead cells, the trypan blue passes through the cell membrane and dead cells appear blue in colour. Cells that had absorbed the trypan blue dye were considered not viable. Routinely only cell cultures exhibiting a satisfyingly high level of viability (>95%) were considered acceptable for experimental use.

A common device used for cell counting is the counting chamber. The most widely used type of chamber is a haemocytometer, and it was originally designed for performing blood cell counts. To prepare the

61

counting chamber the mirror-like polished surface was carefully cleaned with lens paper. The cover slip was also cleaned. Cover slips for counting chambers were specially made and are thicker than those for conventional microscopy, since they must be heavy enough to overcome the surface tension of a drop of liquid. The main divisions separate the grid into 9 large squares (like a tic-tac-toe grid) as shown in Figure 3.1. Each square has a surface area of one square mm, and the depth of the chamber is 0.1 mm. Cell suspensions were mixed appropriately so that the cells did not overlap each other on the grid, and were uniformly distributed.

To determine the cells count, an aliquot of 20 µL of cell suspension after the centrifugation as described in Section 3.4.3 was mixed with 0.4% (v/v) trypan blue solution at 50:50 ratio and left for one min to allow the penetration of the dye to any non-viable cells. Then a pipette was used to transfer 10 µL of the cell suspension mixed with trypan blue dye to the edge of the cover slip on the haemocytometer. The cells were counted using a hand tally counter under an optical microscopy at 10 x objective and focus on one set of 16 corner square (Figure 3.1). The haemocytometer was moved to another three sets of 16 corner squares to carry on counting.

62 Figure 3. 1 A layout of the Neubeur haemocytometer under the microscopy. There are nine 1.0 mm² x 1.0 mm² large squares isolated from each other by the triple lines. The total cell count were taken from area A, B, C and D with each of the larger corner 1mm² consisting 16 small squares.

The total cell counting numbers from all 4 sets of 16 corner squares = (cells/mL x 10⁴) x4 squares from each haemocytometer grid. The overall cell count is divided by 4 to get a mean of each 16 squares and multiply by 2 to adjust for the dilution factor to get the final cell count. For example, if the total cell count from all four sets of 16 corner squares is 100, then the final cell density is 100/4 x 2 x 10⁴/mL.

63

3.4.5 Long term cell storage

The cell pellet after centrifugation was re-suspended in cryopreservation medium (DMEM with 10% FCS and 10% DMSO). The cell suspension was divided into 1ml cryopreservation vials in the concentration of 3.75 x 10⁵ cells/mL. All the vials were labelled with cell line name, passage number, concentration and UK). The Alamar Blue assay incorporates a fluorometric/colorimetric growth indicator based on the detection of metabolic activity. The system incorporates an oxidation-reduction (REDOX) indicator that both fluoresces and changes colour in response to a chemical reduction of growth medium resulting from cell growth. As cells grow in culture, innate metabolic activity results in a chemical reduction of the immediate surrounding environment. Continued growth maintains a reduced environment while inhibition of growth maintains an oxidized environment. Reduction related to growth causes the REDOX indicator to change from oxidised resazurin (non-fluorescent, blue) to a reduced form resorufin (fluorescent, red) as shown in Figure 3.2. In this study, the Alamar blue assay was mainly used to monitor cell proliferation in the continuous culture systems and to further explore the proliferation trend within specific time period in a range of culture conditions.

O

Figure 3. 2 The conversion of resazurin to resorufin. Resazurin is a non-fluorescent compound, which is converted to highly red fluorescent resorufin via reduction reactions of living cells. There is a proportional relationship between the amount of fluorescence produced and the total number of living cells.

64

In general, Alamar blue stock solution was diluted into fresh cell growth medium to get 10% working solution. This working solution was mixed and sterilised with 0.2 µm filters and then kept in the fridge and wrapped in foil until ready to use. To assess cell standard viability over a period of time, triplicate samples were required for each measurement. HUVECs were allowed to grow on cell culture plastics until 80% confluent, and cells were harvested, counted and diluted to gain a range of cell concentrations of 5.0 x 10², 1 x 10³, 1 x 10⁴ and 1 x 10⁵ cells/mL in growth media. A volume of 100 μL of cell suspension was added to each assay well of a 96 well plate and incubated at 37^oC with 5% CO2 to allow cells to attach to the surface. The next morning, the media was aspirated from the wells and cells were washed three times with sterile PBS. 100 μL of Alamar Blue working solution was added into each well and incubated at 37^oC with 5% CO2 for 90 min and the plate was wrapped in foil to protect from the light.

Then the plate was read with a spectrometer at 570nm and the reading was considered as t=0 hr. Further reading were taken at 1, 6, 12, 24, 48 and 72 hr. Empty wells with no cells attached were used as blank.