Three-Dimensional (3D) in vitro cell culture protocols to ...

The part of the protocol described in this peer-reviewed article &#;U-251MG Spheroid generation using low attachment plate method&#; is published on protocols.io, https://dx.doi.org/10./protocols.io.bszmnf46 and is included for printing as S1 File with this article. The part of the protocol described in this peer-reviewed article &#;U-251MG Spheroid Generation Using Hanging Drop Method Protocol&#; is published on protocols.io, https://dx.doi.org/10./protocols.io.btstnnen and is included for printing as S2 File with this article. The part of the protocol described in this peer-reviewed article &#;U-251MG Spheroid generation using a scaffold based method protocol&#; https://dx.doi.org/10./protocols.io.bszqnf5w is published on protocols.io, and is included for printing as S3 File with this article.

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The research project was approved by TU Dublin Research Ethics and Integrity Committee and involved the use of human samples. Human cancer cell lines (U-251 MG, U87 MG and A-172) were used in the study. These are cell lines obtained from reputable commercial cell banks, these are established, commercially available cell lines and consent was not obtained from the original donors. Animal tissue (fetal calf serum) was also used in the study. This was obtained from a reputable commercial company.

The human glioblastoma multiforme cell line (U&#;251 MG, formerly known as U&#;373MG&#;CD14) was a gift from Michael Carty (Trinity College Dublin), U87 MG and A-172 human glioblastomas were purchased from an ATCC European Distributor (LGC Standards). The absence of mycoplasma was checked by using a MycoAlert PLUS Mycoplasma Detection kit (Lonza). Cells were maintained in Dulbecco&#;s modified Eagle medium (DMEM)&#;high glucose supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin. Cells were maintained in a humidified incubator containing 5% CO2 atmosphere at 37°C in a TC flask T25, standard for adherent cells (Sarstedt). Cells were routinely sub-cultured when 80% confluence was reached using 0.25% w/v Trypsin&#;EDTA solution.

U&#;251 MG, U-87 MG and A-172 human glioblastoma cells were used to generate tumour spheroids. Separately, the single cell suspensions were centrifuged at rpm for 5 min, removed the supernatant, tapped the tube and re-suspended the cell pellet in DMEM&#;high glucose supplemented with 10% FBS and 1% penicillin/streptomycin. The single&#;cell suspensions (with desired seeding density) were transferred to a sterile reservoir and seeded 200 μl/well into Nunclon&#; Sphera&#; 96&#;well low attachment plates (Thermo Fisher Scientific) using a multichannel pipette ensuring pipette tips do not touch the surface of the wells to protect the surface coating. The low attachment plates were centrifuged at rpm for 5 min followed by incubation (37°C, 5% CO2, 95% humidity). After 24h of incubation, the media must be replenished. 100 μl media was removed without disrupting the tumourspheres and 100 μl of fresh media (DMEM + 10%FBS + 1%Ab) was added into each well and incubate at 37°C (5% CO2, 95% humidity). The sides of wells should be used to remove or add media, and pipetting should be carried out at average or below average speeds to avoid disruption to spheroids. Tumour spheroid formation was observed within 4 days for U&#;251 MG, U-87 MG and A-172. Tumour spheroid formation was visually confirmed daily using an Optika XDS&#;2 trinocular inverse microscope equipped with a Camera ISH500, and their mean diameters were analyzed using &#;ImageJ version 1.53.e&#; software.

U-251 MG, U-87 MG and A-172 single cell suspensions (with desired seeding densities) were used to generate tumour spheroids using HDP Perfecta3D® 96-Well Hanging Drop Plate. Sterile PBS was added to the reservoirs located on the peripheral rims, which are divided into sections. 2 ml of PBS was added to each plate reservoir section, and 1ml was added per tray reservoir section. This prevented the hanging drop from drying throughout the incubation period. In order to achieve cells per 20 μl of hanging drop, the single cell suspensions prepared in DMEM&#;high glucose supplemented with 10% FBS and 1% penicillin/streptomycin at a concentration of 2.5x10 5 cells/ml. Each hanging drop well was able to hold 20&#;50 μl of cell suspension, and any volume above 50 μl resulted in droplet instability. Hanging drops can be formed by carefully pipetting 20&#;50 μl of cell suspension into the centre of each well from the top side of the plate. Hanging drops should be formed on and confined to the bottom of the plate. Placed the lid on the plate and inserted the assembly into a humidified cell culture incubator at 37°C and 5% CO 2 . Tumour spheroids formation was visually confirmed within 4 days for U&#;251 MG, U-87 MG and A-172. 5μl of fresh media was added back into the hanging drops by placing the end of the pipette tips in the neck region of the access holes/wells and the fresh media was slowly dispensed into the access holes. Once formed, tumourspheres can be transferred from the hanging drop plate to low attachment plates/pre-coated wells in the dish by adding 50 μl of fresh media into each hole.

U-251 MG, U-87 MG and A-172 single cell suspensions (with desired seeding densities) were used to generate tumour spheroids using cellusponge 3D scaffolds. A 9 mm cellusponge disk was slowly placed in the middle of each well in a 24-well plate and 100 μL was seeded from a cell suspension with a cell density of k cells/ml. Cellusponge disks with cells were incubated at 37°C (5% CO 2 , 95% humidity) for 3 hours to remove any air bubbles within the cellusponge. After incubation, 500 μL of DMEM&#;high glucose supplemented with 10% FBS and 1% penicillin/streptomycin was added slowly along the edge of each well in a 24-well plate. Plates with cellusponge scaffolds were incubated overnight at 37°C, 5% CO 2 , 95% humidity. After overnight incubation, the seeded scaffolds were transferred into a new well plate, the media was replenished and the culture should be continued. Tumour spheroid formation was observed within 3&#;4 days.

Tumour spheroid formation was visually confirmed daily using an Optika XDS-2 trinocular inverse microscope equipped with a Camera ISH500, and their mean diameters were analysed using &#;ImageJ version 1.53.e&#; software ( http://imagej.nih.gov/ij/ ). ImageJ is a free software that can be used for manually counting the cell numbers and calculating the cellular size (area / diameter). The ImageJ program was calibrated (set scale) using an image obtained from the same microscope with a known scale before it was used to calculate the cell size (in diameter). Following the calibration, the pictures of the tumourspheres were opened in the program, and a line was drawn across the diameter to measure the tumoursphere&#;s size. The diameters of the spheroids were measured at least three times to obtain the mean and standard deviation.

The growth of U-251 MG, U-87 MG and A-172 tumourspheres were analysed during different incubations (ranging from 24 to 168 h). Cells (initial seeding density was cells/ml) were seeded in the above mentioned Nunclon&#;Sphera&#;96&#;well&#;low attachment plates. Fresh media were added every third day by replenishing old media in each well without disturbing the tumourspheroids. In the hanging drop plate method, cells/well were seeded in the HDP Perfecta3D® 96-well Plate. While in scaffold based method, k cells/ml were seeded in the hydroxipropylcellulose scaffold. The spheroid formation and growth were monitored daily by using an inverted phase-contrast microscope, and the sizes of the spheroids were measured as described above using at least nine spheroids within the three biological repetitions.

For growth analysis, varying numbers of U-251 MG, U-87 MG and A-172 cells (ranging from to 40 000 cells/ml) were seeded in the above mentioned Nunclon&#; Sphera&#; 96-well-low attachment plates for 96 hours. Fresh media were added every third day by replenishing old media in each well without disturbing the tumourspheroids. In the hanging drop plate method, U-251 MG cells (ranging from to cells/well) were seeded in the above mentioned HDP Perfecta3D® 96-well Plate. While in scaffold based method, varying numbers of U-251 MG cells (ranging from 1x10 6 to 6x10 6 cells/ml) were seeded in the hydroxipropylcellulose scaffold. The spheroid formation was monitored after 96 h by using an inverted phase-contrast microscope, and the sizes of the spheroids were measured as explained above, using at least nine spheroids within the three independent experiments.

Spheroid cell health was analysed using the Alamar Blue&#; cell viability reagent (Thermo Fisher Scientific). After the post treatment incubation, tumourspheres were washed with sterile phosphate-buffered saline (PBS), trypsinized using a 0.25% w/v trypsin&#;EDTA solution and incubated for 3 h at 37°C with a 10% Alamar Blue&#; solution [ 10 ]. During the scaffold based method tumourspheres embedded in the cellusponge 3D scaffolds were incubated for 24h instead of 3h [ 24 ]. Fluorescence was measured using an excitation wavelength of 530 nm and an emission wavelength of 590 nm with a Varioskan Lux multiplate reader (Thermo Scientific). The fluorescence signals were normalized by spheroid size (in diameter); a higher ratio indicates healthier spheroids. The experiments consisted of three independent tests in which at least nine spheroids were measured throughout three biological repeats.

3D cell viability was analysed using the CellTiter-Glo® 3D Cell viability assay (Promega). After the post treatment incubation, homogeneous tumourspheres were removed from the 96-well low attachment culture plate and placed separately in single wells of a 96-well plate (Sarstedt). CellTiter-Glo® 3D reagent was added to each well and the luminescence signals were read after 25 minutes of incubation at room temperature using the Varioskan Lux multiplate reader (Thermo Scientific).

Dose response curves for the commonly employed chemotherapeutic drug, Temozolomide (TMZ), used for the treatment of U-251 MG, U-87 MG, and A-172 GBM tumour spheroids. TMZ was dissolved in dimethyl sulfoxide (DMSO) and stored at &#;20°C. These stocks were subsequently used to make the working standard solutions in media. The highest concentration of DMSO used was 0.5%. U-251 MG, U-87 MG and A-172 cells were seeded at a density of 1 × 10 4 cells/ml (200 μl culture medium/well) into Nunclon&#; Sphera&#; 96&#;well low attachment plates (Thermo Fisher Scientific). After tumoursphere construction, existing media were removed from each well and tumourspheres were treated with TMZ (concentration gradient from 500 μM to 0.97 μM), and incubated for 6 days at 37°C (5% CO 2 , 95% humidity). DMSO (20%) was used as a positive control. After the post treatment incubation, the cytotoxicity of the tumourspheres were measured using the both CellTiter-Glo® 3D Cell viability assay and Alamar Blue&#; cell viability reagents as mentioned above, using at least three independent tests with a minimum of three replicates per experiment.

All the experiments were replicated at least three independent times. Prism versions 9.1.0, GraphPad Softwares, Inc. were used to carry out curve fitting and statistical analysis. Dose&#;response curves were measured using nonlinear regression. Data are presented as a percentage and error bars of all figures were presented using the standard error of the mean (SEM), multiple comparison analyzes were performed using Tukey&#;s test unless otherwise stated.

Expected results and discussion

GBM is distinguished by increased vascularization, significant cell heterogeneity, self-renewing cancer stem cells and the interactions between tumour and microenvironment, all of which contribute to tumour progression [25]. Tumour development, metastasis, angiogenesis, cytotoxicity resistance, and immune cell modulation are all influenced by the tumour microenvironment (TME) [10,26]. There is a gap in mostly accessible GBM pre-clinical models and 3D cell culture is able to fill this gap by providing more reliable models to study the correlation between TME, tumour reoccurrence and therapy resistance.

Three distinct approaches, such as low attachment plate ( ), (S1 File), hanging drop plate ( ), (S2 File), and scaffold based methods ( ), (S3 File) were used to create U-251MG, U-87 MG and A-172 3D human glioblastoma cell culture models. This facilitated 3D cell&#;cell and cell&#;ECM interactions and mirrored the diffusion-limited distribution of oxygen, nutrients, metabolites, and signaling molecules seen in the microenvironment of in vivo tumours [10]. Most research to date has used 2D cell culture ( ), which has limitations as experimental models to predict biological responses, as explained previously.

U-251 MG, U-87 MG, and A-172 human glioblastoma astrocytoma spheroids formation and growth were monitored daily by using an inverted phase&#;contrast microscope, and their mean diameters were analysed using &#;ImageJ version1.53.e&#; software for at least three independent experiments. U-251MG tumoursphere growth during the low attachment plate method was found to be significantly increased with the incubation time, the size ranging from 135 μm, 229 μm, 323 μm and 461 μm ( ) for 24 to 96 h incubation respectively. U-87 MG tumourspheres were significantly increased with the incubation time, the size range from 129 μm, 234 μm, 303 μm and 357 μm ( ) for 24 to 96 h incubation respectively. While, A-172 tumourspheres also showed same behaviour with the increasing incubation time and the sizes rage from 71 μm, 191 μm, 240 μm and 367 μm ( ) for 24 to 96 h incubation respectively.

U-251 MG tumoursphere growth during the hanging drop plate method was shown to be considerably enhanced with the incubation time, the size ranging from 105 μm, 139 μm, 208 μm and 269 μm ( ) for 24 to 96 h incubation respectively. U-87 MG tumourspheres were significantly increased with the incubation time, the size range from 92 μm, 143 μm, 224 μm and 252 μm ( ) for 24 to 96 h incubation respectively. While, A-172 tumourspheres also showed same behaviour with the increasing incubation time and the sizes rage from 63 μm, 131 μm, 207 μm and 265 μm ( ) for 24 to 96 h incubation, respectively.

U-251 MG tumoursphere growth in hydroxipropylcellulose 3D scaffold was shown to be considerably enhanced with incubation time, with sizes ranging from 22 μm, 49 μm, 70 μm, and 110 μm for 24 to 96 h incubation, respectively ( ). U-87 MG tumourspheres were significantly increased with the incubation time, the size range from 28 μm, 55 μm, 90 μm and 143 μm ( ) for 24 to 96 h incubation respectively. While, A-172 tumourspheres size also significantly enhanced with the increasing incubation time and the sizes rage from 17 μm, 51 μm, 69 μm and 97 μm ( ) for 24 to 96 h incubation respectively.

These results proved that these protocols have the ability to develop 3D tumourspheres and the presence of heterogeneous cellular subpopulations such as actively proliferating, quiescent, hypoxic, and necrotic cells [2,14,27].

The optimum U-251 MG ( ), U-87 MG ( ) and A-172 ( ) tumourspheroids formations were observed within 96 h of incubation for the cells/ml initial seeding density. One&#;way analysis of variance (ANOVA) demonstrated that there were significant differences in tumourspheres diameter during 24&#;96 h incubation, while there was no significant difference during 96&#;168 h incubation. It was also observed that exponential growth (Log) was achieved within the initial 4 days of growth, after which the growth curve became stationary in all these three glioblastoma cell lines.

For growth analysis, varying numbers of U-251 MG, U-87 MG and A-172 cells (ranging from to 40 000 cells/ml) were seeded in the Nunclon&#; Sphera&#; 96&#;well low attachment plates as explained above. The largest U&#;251 MG tumourspheres were observed with 10 000, 15 000, and 20 000 cells/ml initial seeding densities after 96 h of incubation. One-way ANOVA demonstrated that there was a significant difference in tumoursphere diameter between each initial seeding densities as shown in . However, there was no significant difference between diameters in 10 000, 15 000, and 20 000 cells/ml seeding densities. The largest U-87 MG tumourspheres were observed with 40 000 cells/ml initial seeding densities after 96 h incubation. One-way ANOVA demonstrated that there was a significant difference in tumoursphere diameter between each of the initial seeding densities as shown in . However, there was no significant difference between diameters at 10 000 and 15 000 cells/ml seeding densities. While, the largest A-172 tumourspheres were observed with 40 000 cells/ml initial seeding densities after 96h incubation. One-way ANOVA demonstrated that there was a significant difference in tumoursphere diameter between each of the initial seeding densities as shown in . Though, there was no significant difference between diameters in 10 000, 15 000, and 20 000 cells/ml seeding densities.

U-251 MG, U-87 MG and A-172 cells health analysed after 96h incubation using Alamar Blue&#; cell viability reagent as explained above and the fluorescence signals were normalized by spheroid size (diameter in μm). A higher ratio suggests that the spheroids are healthier. During U-251 MG growth confirmed that and 10 000 cells/ml initial seeding densities were having highest spheroids cell health. One-way ANOVA confirmed that there was no significant difference in tumoursphere health during and 10 000 cells/ml ( ). During U-87 MG growth, it was confirmed that 10 000 and 15 000 cells/ml initial seeding densities were having highest spheroids cell health. One-way ANOVA confirmed that there was no significant difference in tumoursphere health during 10 000 and 15 000 cells/ml ( ). During A-172 growth confirmed that 10 000, 15 000 and 20 000 cells/ml initial seeding densities were the ones having the highest spheroids cell health. One-way ANOVA confirmed that there was no significant difference in tumoursphere health during 10 000, 15 000 and 20 000 cells/ml ( ).

We studied tumoursphere growth in low attachment plates with various seeding densities and observed tumoursphere growth ranging in diameter from 150 to 650 μm. Our findings are correlate with the tumoursphere diameters determined by Singh et al [28]. According to the results, cells/ml initial seeding density was the most suitable seeding density for low attachment plate method and all the above glioblastoma cell lines were able to produce healthy tumourspheres after 96h incubation. Recently, we used the same protocol to generate U-251MG tumourspheres and successfully studied the plasma induced cytotoxicity in 3D glioblastoma tumour spheroids [10].

This results also proved low attachment plate&#;s ability to promote aggregation of cells by cell-cell and cell-ECM interactions while blocking the ECM interaction to the plastic surface. Which can be used as a pre-clinical model due to its simplicity, efficiency, higher reproducibility and also possible to generate a wide range of tumour cell types using the same protocol in any laboratory conditions [9,15,16,29].

The optimum U-251 MG ( ), U-87 MG ( ) and A-172 ( ) tumourspheroids formations were attained after 96 h of incubation for the cells/well initial seeding density by achieving a size range of 251&#;285 μm, 252&#;279 μm and 217&#;265 μm respectively. One&#;way ANOVA indicated that there were significant differences in tumourspheres diameter during 24&#;96 h incubation, while, there were no significant difference during 96&#;168 h incubation. It was also observed that exponential growth (Log) was achieved within the initial 4 days of growth, after which the growth curve became stationary in all these three glioblastoma cell lines.

For growth analysis, varying numbers of U&#;251 MG, U-87 MG and A-172 cells (ranging from to 10 000 cells/well) were seeded in the HDP Perfecta3D® 96-well hanging drop plates and the mean sizes were computed after 96h of incubation. The largest U&#;251 MG tumourspheres were observed with 10 000 cells/well initial seeding densities after 96 h incubation. As illustrated in , one-way ANOVA revealed a significant difference in tumoursphere diameter between each initial seeding density. The largest U-87 MG tumourspheres were observed with 8 000 to 10 000 cells/well initial seeding densities after 96 h of incubation. One-way ANOVA demonstrated that there is a significant difference in tumoursphere diameter between each of the initial seeding densities as shown in . However, there was no significant difference between diameters in 5 000, 8 000 and 10 000 cells/well seeding densities. While the largest A-172 tumourspheres were observed with 8 000 to 10 000 cells/well initial seeding densities after 96 h of incubation. One-way ANOVA demonstrated that there is a significant difference in tumoursphere diameter between each of the initial seeding densities as shown in . Though, there was no significant difference between diameters in 4 000 to 5 000, and 8 000 to 10 000 cells/well seeding densities.

During the U-251 MG spheroids cell health investigation, it was established that the initial seeding density of cells/well had the best spheroids cell health. The substantial difference in to cells/well and to cells/well was verified by one-way ANOVA, however there was no significant difference in tumoursphere health at the other seeding densities ( ).

U-87 MG growth confirmed that 5 000 cells/well initial seeding density was having highest spheroids cell health. One-way ANOVA confirmed that there was no significant difference in tumoursphere health during 4 000, 5 000, 8 000 and 10 000 cells/well ( ). During A-172 growth confirmed that 5 000, 8 000 and 10 000 cells/well initial seeding densities were having highest spheroids cell health. One-way ANOVA confirmed that there was no significant difference in tumoursphere health during 5 000, 8 000 and 10 000 cells/well ( ).

We studied tumoursphere growth in hanging drop plates with various seeding densities and observed tumoursphere growth ranging in diameter from 100 to 400 μm. According to the results, cells/well initial seeding density was the most suitable seeding density for the hanging drop plate method, and all the above glioblastoma cell lines were able to produce healthy tumourspheres after 96 h incubation. This result also proved the hanging drop plate&#;s ability to produce uniform sized spheroids, ability to control the size of spheroid by seeding density, higher replicability, lower cost, and ability of tumoursphere mass production within a shorter time period [9,12,15,29].

The largest U-251 MG and A-172 tumourspheroids formation were attained after 120 h incubation by achieving a size range 110&#;156 μm ( ) and 146&#;174 μm ( ) for the k cells/ml initial seeding density respectively. One-way ANOVA indicated that there were significant difference in tumoursphere diameter throughout the incubation. The optimum U-87 MG tumourspheroid formation was observed within 120 h of incubation (size range from 133&#;191 μm) for the k cells/ml initial seeding density. One&#;way ANOVA indicated that there were significant differences in tumourspheres diameter during 24&#;120 h incubation, while, there was no significant difference during 48&#;72 h incubation ( ).

For growth analysis, varying numbers of U&#;251 MG, U-87 MG and A-172 (ranging from 1x106 to 6x106 cells/ml) were seeded in the hydroxipropylcellulose 3D scaffolds. Fresh media were added every third day by replenishing old media in each well without disturbing the scaffolds, and the mean sizes were calculated after 120 h of incubation. The largest U-251 MG tumourspheres were detected with 5x106 and 6x106 cells/ml initial seeding densities after 120 h of incubation. One-way ANOVA verified that there is a significant difference in tumoursphere diameter between 4x106 and 5x106 seeding densities, while there was no significant difference in diameters between 5x106 and 6x106 cells/ml initial seeding densities as shown in . The largest U-87 MG and A-172 tumourspheres were observed with 5x106 and 6x106 cells/ml initial seeding densities after 120 h of incubation. One-way ANOVA demonstrated that there were significant difference in tumoursphere diameter between each initial seeding densities as shown in respectively. However, there was no significant difference between diameters in 5x106 and 6x106 cells/ml seeding densities.

U-251 MG, U-87 MG and A-172 spheroids cell health were analysed after 120 h of incubation as explained above, U-251 MG growth confirmed that 5x106 cells/ml initial seeding density was having highest spheroids cell health. One-way ANOVA confirmed that there were significant difference in tumoursphere health during 4x106, 5x106 and 6x106 cells/ml. While there was a significant difference between 3x106 and 4x106 densities as shown in .

During U-87 MG growth, it was confirmed that 5x106 cells/ml initial seeding densities were having the highest spheroids cell health. One&#;way ANOVA confirmed that there were significant differences in tumourspheres health from 1x106 to 6x106 cells/ml ( ). During A-172 growth, it was confirmed that 5x106 and 6x106 cells/ml initial seeding densities had the highest spheroids cell health. One-way ANOVA confirmed that there was no significant difference in tumoursphere health during 4x106, 5x106 and 6x106 cells/ml ( ).

We studied tumoursphere growth in hydroxipropylcellulose 3D scaffolds with various seeding densities and observed tumoursphere growth ranging in diameter from 50 to 200 μm. According to the results, 5x106 cells/well initial seeding density was the most suitable seeding density for the scaffold based method and all the above glioblastoma cell lines were able to produce healthy tumourspheres after 120 h of incubation. These results also proved the biological scaffold&#;s higher biocompatibility and its possibility to control scaffold&#;s composition, porosity, and elasticity to get better GBM ECM representation [15,20]. This protocol can be applied to hydroxipropylcellulose scaffolds or any other natural scaffold [30] and possible to generate wide range of tumoursphere types in any laboratory. It is also possible to improve scaffold chemistry and composition to mimic the physiological architecture of any glioblastoma tumours.

The effects of TMZ cytotoxicity on the different GBM cell lines were studied using U&#;251 MG, U-87 MG and A-172 tumourspheres. TMZ induced cytotoxicity was studied using two different cell viability assays as shown in . The CellTiter-Glo® 3D Cell viability assay quantifies the amount of ATP present, which is a marker for the presence of metabolically active cells, to determine the number of viable cells in a 3D cell culture. While alamarBlue cell viability, Resazurin is used as an oxidation-reduction (REDOX) indicator that undergoes colorimetric change in response to cellular metabolic reduction.

TMZ treated tumourspheres (concentration gradient from 500 μM to 0.97 μM), were post incubated for 6 days at 37°C. An IC50 of 143.6 μM (135.2 ± 152.6 μM), 71.25 μM (66.41 ± 76.44 μM), and 111.0 μM (103.1 ± 119.4 μM) were found for U&#;251 MG, U-87 MG and A-172 tumourspheres respectively, when analysed by using the Alamar Blue&#; cell viability assay ( ). An IC50 of 174.4 μM (160.5 ± 189.5 μM), 76.06 μM (70.81 ± 81.70 μM), and 134.0 μM (115.5 ± 155.3 μM) were found for U&#;251 MG, U-87 MG and A-172 tumourspheres respectively, when analysed by using the CellTiter-Glo® 3D Cell viability assay ( ). Two&#;way ANOVA demonstrated that there were significant differences in viability between the different TMZ concentrations and different cell lines (p< 0.). According to these results, it was postulated that as U-87 MG has the highest TMZ sensitivity, while U-251 MG tumourspheres showed highest cell viability with TMZ treatment. TMZ induced cytotoxicity in all three cell lines showed similar values, when comparing both cell viability assays. However, comparatively higher IC50 values were observed from the CellTiter-Glo® 3D Cell viability assay, and this may be due to the difference in assay chemistries and metabolic targets in viability assays.

The effects of diffusion of the active dyes through the matrices and their subsequent bioavailability to the cells can lead to misinterpretation of the results obtained. The concern is addressed in the present study by converting tumourspheres into single cells before cell viability analysis using the Alamar Blue&#; cell viability assay. This method can be successfully applied to tumourspheres constructed using low attachment plate and hanging drop plate methods since it is possible to collect cells after the growth / treatment. Bonnier and colleagues reported the way to use the Alamar Blue&#; cell viability assay for tumourspheres constructed using hydrogels or scaffold based methods. During this method tumoursphere embedded in gels were incubated with Alamar Blue&#; for 24h instead of 3h to get high diffusion of the active dyes through the matrices to cells, similar to our study [24]. On the other hand, the CellTiter-Glo® 3D Cell viability assay is quicker, easier to use and directly applies to the tumourspheres constructed using low attachment plate, hanging drop plate method and scaffold based method.

Ultimately, these basic 3D cell culture models can be further improved to study the role of the blood brain barrier and chemotherapeutic resistance in glioblastoma [31], exploring GBM / normal tissue interactions. The potential impact of the microbiome, TME, vasculature, infiltrating parenchymal and peripheral immune cells on glioblastoma treatment techniques can also be further investigated with more advanced 3D co-culture models [32]. In the future, advances in 3D cell culture will make it possible to generate whole 3D in vitro GB organoids, leading to personalized treatments for glioblastoma [20,33,34].

Although there are different classifications for cytotoxicity and cell viability assays, in this chapter, these assays are classified according to measurement types of end points (color changes, fluorescence, luminescent etc.).

Erythrosine B, also known as erythrosine or Red No. 3, is primarily used as food coloring agent [ 20 , 21 ]. Erythrosine B has already been introduced as a vital dye for counting viable cells. Principle of this dye exclusion assay is similar to trypan blue dye exclusion assay principle. Although erythrosine B is an alternative bio-safe vital dye for cell counting; it is not widely used to count viable or dead cells.

While the staining procedure is quite simple, it is difficult to process large number of samples concurrently, particularly where the exact timing of progressive cytotoxic effects is required [ 4 ]. Furthermore, trypan blue staining cannot be used to distinguish between the healthy cells and the cells that are alive but losing cell functions. Therefore, it is not sufficiently sensitive to use for in vitro cytotoxicity testing. Another disadvantage of trypan blue is toxic side effect of this dye on mammalian cells [ 20 ].

For more information, please visit 48 well cell culture plates.

Disadvantages: Cell counting is generally done using a hemacytometer [ 19 ]. Therefore, counting errors (~10%) could be occurred. Counting errors have been attributed to poor dispersion of cells, cell loss during cell dispersion, inaccurate dilution of cells, improper filling of the chamber and presence of air bubbles in the chamber [ 17 ].

Advantages: This method is simple, inexpensive, and a good indicator of membrane integrity [ 17 ], and dead cells are colored blue within seconds of exposure to the dye [ 18 ].

This dye exclusion assay is used to determine the number of viable and/or dead cells in a cell suspension. Trypan blue is a large negatively charged molecule. Trypan blue dye exclusion assay is based on the principle that live cells possess intact cell membranes that exclude this dye, whereas dead cells do not. In this assay, adherent or nonadherent cells are incubated with serial dilutions of test compounds for various times. After the compound treatment, cells are washed and suspended. Cell suspension is mixed with dye and then visually examined to determine whether cells take up or exclude dye. Viable cells will have a clear cytoplasm, whereas dead cells will have a blue cytoplasm [ 14 , 15 ]. Number of viable and/or dead cells per unit volume is determined by light microscopy as a percentage of untreated control cells [ 15 , 16 ].

Dye exclusion assays have unique advantages for chemosensitivity testing. They are comparatively simple, require small numbers of cells, are rapid, and are capable of detecting cell kill in nondividing cell populations. Further investigations into the possible role of these assays in chemosensitivity testing are warranted [ 11 ]. However, none of these dyes is recommended for use on monolayer cell cultures but rather they are intended for cells in suspension; thus monolayer cells must first trypsinized [ 6 ].

If dye exclusion assays are used, following factors must be considered (i) lethally damaged cells by cytotoxic agents may require several days to lose their membrane integrity, (ii) the surviving cells may continue to proliferate during this time, and (iii) some lethally damaged cells are not appear to be stained with dye at the end of the culture period, because they may undergo an early disintegration. Factors (ii) and (iii) may cause an underestimate of cell death when the results of the assay are based on percent viability expression [ 11 , 12 , 13 ].

The proportion of viable cells in a cell population can be estimated in various methods. The simplest and widely used one of the methods is dye exclusion method. In dye exclusion method, viable cells exclude dyes, but dead cells not exclude them. Although the staining procedure is quite simple, experimental procedure of large number of samples is difficult and time consuming [ 4 ]. Determination of membrane integrity is possible via dye exclusion method. A variety of such dyes have been employed, including eosin, Congo red, erythrosine B, and trypan blue [ 5 , 6 ]. Of the dyes listed, trypan blue has been used the most extensively [ 7 , 8 , 9 , 10 ].

2.2. Colorimetric assays

Principle of colorimetric assays is the measurement of a biochemical marker to evaluate metabolic activity of the cells. Reagents used in colorimetric assays develop a color in response to the viability of cells, allowing the colorimetric measurement of cell viability via spectrophotometer. Colorimetric assays are applicable for adherent or suspended cell lines, easy to perform, and comparably economical [22, 23]. Commercial kits of colorimetric assays are available from several companies and generally experimental procedures of these assays are available in kit packages.

2.2.1. MTT assay

MTT (3-(4,5-dimethylthiazol-2-yl)-2&#;5-diphenyltetrazolium bromide) assay is one of the most commonly used colorimeteric assay to assess cytotoxicity or cell viability [24]. This assay determines principally cell viability through determination of mitochondrial function of cells by measuring activity of mitochondrial enzymes such as succinate dehydrogenase [18]. In this assay, MTT is reduced to a purple formazan by NADH. This product can be quantified by light absorbance at a specific wavelength.

Advantages: This method is far superior to the previously mentioned dye exclusion methods because it is easy to use, safe, has a high reproducibility, and is widely used to determine both cell viability and cytotoxicity tests [18, 25].

Disadvantages: MTT formazan is insoluble in water, and it forms purple needle-shaped crystals in the cells. Therefore, prior to measuring the absorbance, an organic solvent such as dimethyl sulfoxide (DMSO) or isopropanol is required to solubilize the crystals. Additionally, the cytotoxicity of MTT formazan makes it difficult to remove cell culture media from the plate wells due to floating cells with MTT formazan needles, giving significant well-to-well error [18, 26].

Additional control experiments should be conducted to reduce false-positive or false-negative results that caused by background interference due to inclusion of particles. This interference could lead to an overestimation of the cell viability. This can often be controlled by subtraction of the background absorbance of the cells in the presence of the particles, but without the assay reagents [18, 26].

2.2.2. MTS assay

The MTS assay (5-(3-carboxymethoxyphenyl)-2-(4,5-dimethyl-thiazoly)-3-(4-sulfophenyl) tetrazolium, inner salt assay) is a colorimetric assay. This assay is based on the conversion of a tetrazolium salt into a colored formazan by mitochondrial activity of living cells. The amount of produced formazan is depend on the viable cell number in culture and can be measured with spectrophotometer at 492 nm.

Advantages: Previous studies suggest that the MTS in vitro cytotoxicity assay combines all features of a good measurement system in terms of ease of use, precision, and rapid indication of toxicity [27, 28]. MTS assay is a rapid, sensitive, economic, and specific in vitro cytotoxicity assay. Performance of this assay is very competitive to other toxicological tests. This assay provides ideal properties for cytotoxicity measurement because it is easy to use, rapid, reliable, and inexpensive. Therefore, it can be used for onsite toxicological assessments [27, 29, 30, 31].

Disadvantages: The level of absorbance measured at 492 nm is influenced by the incubation time, cell type, and cell number. The proportion of MTS detection reagents to cells in culture also influences the measured absorbance level. Previous studies suggested a linear relationship between incubation time and absorbance for short incubation times up to 5 hours [29, 32, 33]. Therefore, proper incubation times for this assay are 1&#;3 hours.

2.2.3. XTT assay

A colorimetric method based on the tetrazolium salt XTT (2,3-bis(2-methoxy-4-nitro-5-sulphophenyl)-5-carboxanilide-2H-tetrazolium, monosodium salt) was first described by Scudiero et al. [34]. While MTT produced a water-insoluble formazan compound which required dissolving the dye in order to measure its absorbance, the XTT produces a water-soluble dye. The procedure of XTT is simply for measuring proliferation and is therefore an excellent solution for quantitating cells and determining their viability. XTT is used to assay cell proliferation as response to different growth factors. It is also used for assaying cytotoxicity.

This assay is based on the ability reduction of the tetrazolium salt XTT to orange-colored formazan compounds by metabolic active cells. Orange-colored formazan is water soluble and its intensity can be measured with a spectrophotometer. There is a linear relationship between the intensity of the formazan and the number of viable cells. The use of multiwell plates and a spectrophotometer (or ELISA reader) allows for study with a large number of samples and obtaining results easily and rapidly. The procedure of this assay includes cell cultivation in a 96-well plate, adding the XTT reagent and incubation for 2&#;24 hours. During the incubation time, an orange color is formed and the intensity of color can be measured with a spectrophotometer [34, 35].

Advantages: XTT assay is speed, sensitive, easy to use, and safe method. It has high sensitivity and accuracy [35].

Disadvantages: XTT assay performance depends on reductive capacity of viable cells with the mitochondrial dehydrogenase activity. Therefore, changes of reductive capacity of viable cells resulting from enzymatic regulation, pH, cellular ion concentration (e.g., sodium, calcium, potassium), cell cycle variation, or other environmental factors may affect the final absorbance reading [34, 35].

2.2.4. WST-1 assay

WST-1 (2-(4-iodophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H tetrazolium monosodium salt) cell proliferation assay is a simple, colorimetric assay designed to measure the relative proliferation rates of cells in culture. The principle of this assay is based on the conversion of the tetrazolium salt WST-1 into a highly water-soluble formazan by mitochondrial dehydrogenase enzymes in the presence of intermediate electron acceptor, such as mPMS (1-methoxy-5-methyl-phenazinium methyl sulfate) [36]. The water-soluble salt is released into the cell culture medium. Within incubation period, the reaction produces a color change which is directly proportional to the amount of mitochondrial dehydrogenase in cell culture and thus, the assay measures the metabolic activity of cells.

To perform the assay, the WST-1 reagent that is ready-to-use is added directly into the media of cells cultured in multiwell plates. The cultures are then given 30 minutes&#;4 hours to reduce the reagent into the dye form. The plate is then immediately read at 450 nm with a reference reading at 630 nm [37].

Advantages: It is easy to use, safe, has a high reproducibility, and is widely used to determine both cell viability and cytotoxicity tests. Furthermore, phenol red indicators in cell culture medium do not interfere with the dye reaction. Because the colored dye which produced at the end of experiment is water-soluble, it is not required a solvent and additional incubation time [37].

Disadvantages: The standard incubation time of WST-1 time is 2 h. Whether one-time addition of WST-1 can reflect the effect of the testing agents at different time points on the trend of relative cell viability is still unclear [37].

2.2.5. WST-8 assay

WST-8 assay is a colorimetric assay for the determination of viable cell numbers and can be used for cell proliferation assays as well as cytotoxicity assays. WST-8 (2-(2-methoxy-4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H tetrazolium, monosodium salt), a highly stable and water-soluble WST, is utilized in Cell Counting Kit-8 (CCK-8). It is more sensitive than WST-1 particularly at neutral pH [37]. Because of the electron mediator, 1-methoxy PMS in this kit is highly stable, and CCK-8 is stable for at least 6 months at the room temperature and for 1 year at 0&#;5oC. Since WST-8, WST-8 formazan, and 1-methoxy PMS have no cytotoxicity on cells in the culture media, same cells from the previous assay may be used for additional experiments.

Advantages: WST-8 is not cell permeable, which results in low cytotoxicity. Therefore, after the assay, it is possible to continue further experiments using the same cells. Furthermore, it produces the water-soluble formazan upon cellular reduction, which would provide an additional advantage to the method by allowing a simpler assay procedure and not required an extra step to dissolve the formazan [28].

Disadvantages: An important consideration is that reduction of assay substrates is impacted by changes in intracellular metabolic activity that has no direct effect on overall cell viability [15].

2.2.6. LDH (lactate dehydrogenase) assay

LDH (lactate dehydrogenase) cytotoxicity assay is a colorimetric method of assaying cellular cytotoxicity. LDH Cytotoxicity Assay Kit can be used with different cell types not only for assaying cell-mediated cytotoxicity but also for assessment of cytotoxicity mediated by toxic chemicals and other test compounds. The assay measures the stable, cytosolic, lactate dehydrogenase (LDH) enzyme quantitatively. This enzyme releases from damaged cells. LDH is an enzyme that is normally found within the cell cytoplasm. When cell viability reduced leakiness of the plasma membrane increase and therefore LDH enzyme is released into the cell culture medium. The released LDH is measured with a coupled enzymatic reaction that results in the conversion of a tetrazolium salt (iodonitrotetrazolium (INT)) into a red color formazan by diaphorase. In the first step, LDH catalyze conversion of lactate to pyruvate and thus NAD is reduced to NADH/H+. In a second step, catalyst (diaphorase) transfers H/H+ from NADH/H+ to the tetrazolium salt 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyltetrazolium chloride (INT), which is reduced to red formazan [38, 39].

The LDH activity is determined as NADH oxidation or INT reduction over a defined time period. The resulting red formazan absorbs maximally at 492 nm and can be measured quantitatively at 490 nm.

The detergent Triton X-100 is commonly used as positive control in the LDH assay to determine the maximum LDH release from the cells. In addition, well-known membranolytic particles such as crystalline silica can be used as a positive control in LDH assay [40].

Advantages: Reliability, speed, and simple evaluation are some of characteristics of this assay. Because, the loss of intracellular LDH and its release into the culture medium is an indicator of irreversible cell death due to cell membrane damage [38, 41].

Disadvantages: The major limitation of this assay is that serum and some other compounds have inherent LDH activity. For example, the fetal calf serum has extremely high background readings. Therefore, this assay is limited to serum-free or low-serum conditions, limiting the assay culture period (depending on your cells&#; tolerance to low serum) and reducing the scope of the assay as it can no longer allow determination of cell death caused under normal growth conditions (i.e. in 10% fetal calf serum). At a minimum, you should always first test the assay with an unused aliquot of the media you intend to use and compare the reading to that from media lacking supplements (e.g. straight DMEM) [42].

2.2.7. SRB (Sulforhodamine B) assay

SRB (Sulforhodamine B) assay is a rapid and sensitive colorimetric method for measuring the drug-induced cytotoxicity in both attached and suspension cell cultures. This assay as first described by Skehan and colleagues was developed for use in the disease-orientated, large-scale anticancer drug discovery program of the National Cancer Institute (NCI) that was launched in . SRB is a bright pink aminoxanthene dye with two sulfonic groups. Under mildly acidic conditions, SRB binds to protein basic amino acid residues in TCA-fixed (trichloroacetic acid) cells to provide a sensitive index of cellular protein. SRB assay is also used to evaluate colony formation and colony extinction [43].

Advantages: The SRB assay is simple, fast, and sensitive. It provided good linearity with cell number, permitted the use of saturating dye concentrations, is less sensitive to environmental fluctuations, is independent of intermediary metabolism, and provided a fixed end point that is not require a time-sensitive measurement of initial reaction velocity [43]. Reproducibility of this assay is high.

Disadvantages: It is important to obtain and maintain a homogeneous cell suspension. Cellular clumps/aggregates should be avoided for high assay performance.

2.2.8. NRU (neutral red uptake) assay

The neutral red uptake (NRU) assay is also one of the most used colorimetric cytotoxicity/cell viability assay. This assay was developed by Borenfreund and Puerner [44]. This assay was based on the ability of viable cells to take up the supravital dye neutral red. This weakly cationic dye penetrates cell membranes by nonionic passive diffusion and concentrates in the lysosomes. The dye is then extracted from the viable cells using an acidified ethanol solution and the absorbance of the dye is measured using spectrophotometer.

Neutral red uptake depends on the capacity of cells to maintain pH gradients through the ATP production. At physiological pH, net charge of the dye is zero. This charge enables the dye to penetrate the cell membranes. Inside the lysosomes, there is a proton gradient to maintain a pH lower than that of the cytoplasm. Thus, the dye becomes charged and is retained inside the lysosomes. When the cell dies or pH gradient is reduced, the dye cannot be retained. In addition, the uptake of neutral red by viable cells can be modified by alterations in cell surface or lysosomal membranes. Thus, it is possible to distinguish between viable, damaged, or dead cells [44]. Lysosomal uptake of neutral red dye is a highly sensitive indicator of cell viability. The assay can quantitate cell viability and measure cell replication, cytostatic effects or cytotoxic effects depending on the seeding density [45]. Absorbance is measured at 540 nm in multiwell plate reader spectrophotometer.

Advantages: NRR assay is a good marker of lysosomal damage. Also, speed and simple evaluation are some advantages of this assay.

Disadvantages: It has been reported that the NRR assay is either minimally or not at all affected by natural factors, such as temperature and salinity, but is mainly influenced by pollutants [46].

2.2.9. CVS assay (crystal violet assay)

Adherent cells detach from cell culture plates during cell death. This feature can be used for the indirect assessment of cell death and to determine differences in proliferation rate upon stimulation with cytotoxic agents. One simple method to detect maintained adherence of cells is crystal violet assay. In this assay, crystal violet dye binds to proteins and DNA of viable cells, and thus, attached cells are stained with this dye. Cells lose their adherence during cell death and are subsequently lost from the population of cells, reducing the amount of crystal violet staining in a culture. Crystal violet assay is a quick and reliable screening method that is suitable for the examination of the impact of chemotherapeutics or other compounds on cell survival and growth inhibition [47].

Advantages: Crystal violet staining is a quick and versatile assay for screening cell viability under diverse stimulation conditions [48]. However, it is potentially compromised by proliferative responses that occur at the same time as cell death responses. Therefore, chemical inhibitors of caspases and/or of necroptosis may be incorporated into the assay [49, 50]. Alternatively, molecular studies (e.g., overexpression or knockdown) can be performed to more specifically address the nature of cell death [51].

Disadvantages: Crystal violet assay is insensitive to changes in cell metabolic activity. Therefore, this assay is not appropriate for studies used cell metabolism affected compounds. While crystal violet assay is suitable for the examination of the impact of chemotherapeutics or other compounds on cell survival and growth inhibition, it is not able to measure cell proliferation rate [51].

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