Photoporation is a rapidly expanding technique for the introduction of macromolecules into single cells. However, there remains no study into the true efficiency of this procedure. Here, we present a detailed analysis of transfection efficiency and cell viability for femtosecond optical transfection using a titanium sapphire laser at 800 nm. Photoporation of 4000 Chinese Hamster ovary cells was performed, representing the largest optical transfection study reported to date. We have investigated a range of laser fluences at the cell membrane and, at 1.2 μJ/cm2, have found an average transfection efficiency of 50 ± 10%. Contrary to recent literature, in which 100% efficiency is claimed, our measure of efficiency accounts for all irradiated cells, including those lost as a result of laser treatment, thereby providing a true biological measure of the technique.
©2006 Optical Society of America
The introduction of membrane impermeable substances such as foreign DNA into a cell (transfection) is a ubiquitous problem in cell biology. This technique is particularly challenging when it is desirable to target specific cells for treatment, as most transfection technologies (such as electroporation and liposomal transfection) are based on treating a global population of cells simultaneously . Laser-assisted cell poration (photoporation) offers this distinct advantage of cell specificity, while maintaining high transfection efficiency, good post-transfection cell viability and overall ease of operation. The optical transfection technique can utilize both continuous-wave [2–5] and pulsed [6–9] laser sources, but the femtosecond titanium sapphire laser [10–12] has recently proven highly successful, mainly due to the superior peak powers of ultrashort pulses. Femtosecond lasers and photoporation have the potential to form the basis for a future biophotonics toolkit incorporating confocal and multiphoton imaging for lab-on-a-chip, single-cell or microfluidic configurations in a fully sterile environment. Despite the current interest in femtosecond optical transfection, very little research has been carried out to investigate the realistic potential of the technique in comparison to established methodologies.
In this paper, we have determined the first full and accurate representation of femtosecond optical transfection efficiency in order to provide a true biological measure for the technique. We account for each and every targeted cell, including those lost as a result of laser treatment, in order to provide a clear understanding of the relative usefulness of optical transfection in a biological environment. This contrasts to previous measures of efficiency in the literature, where those cells killed as a result of the photoporation process have typically been excluded.
We perform photoporation of four thousand Chinese Hamster ovary (CHO) cells using a highly-focused femtosecond titanium sapphire laser in a home-built inverted optical microscope. We also measure the characteristic cell viability as a result of the laser procedure. CHO cells have long been used by cell biologists as an experimental mammalian model system, and in the last few years they have been a common cell type used to illustrate the potential of optical transfection. We investigate a wide range of laser fluences at the cellular membrane to determine the optimum femtosecond laser parameters for optimum CHO cell transfection efficiency. In addition, we perform photoporation of cells in the presence of a membrane-impermeable dye which provides conclusive evidence of the cell membrane being permeabilised by the action of the focused laser beam. Finally we observe that the size and duration of generated holes in photoporated cells can be used to determine the likelihood of cellular viability following laser poration.
2. Photoporation and transfection apparatus
In the technique of photoporation, a laser beam is typically focused through a high numerical aperture microscope objective lens onto the plasma membrane of the targeted cell. The brief presence of the highly localized laser beam serves at least to modify the permeability of the cell membrane, if not creating a transient pore, thereby allowing foreign DNA in the surrounding medium to enter before the pore closes, and the cell heals itself. If the foreign DNA is then transcribed and translated into protein, then optical transfection has occurred. If another membrane impermeable macromolecule is introduced inside the cell, phototranslocation has occurred. When utilizing a tightly-focused femtosecond near-infrared laser, the photoporation mechanism relies heavily on a two-photon process , and therefore provides an inherently high level of focusing precision in the sub-micron region. In addition, the nanosecond  and femtosecond  laser can be employed to photoporate cells in suspension as well as those which are adherent to a surface. Interestingly, we have recently shown that the equivalent single-photon approach can also be effective, as demonstrated with a relatively inexpensive continuous-wave violet diode laser operating around 400 nm .
For our investigations, we made use of an inverted optical microscope configuration (Fig. 1) with a femtosecond titanium sapphire laser (800 nm, 80 MHz, 120 fs, 1.5 W output). This laser provided a high-quality circularized Gaussian beam (dx = dy) with an associated M-squared parameter of <1.2 in both transverse planes. The laser output was first passed though a variable neutral density (ND) filter to attenuate the beam-focus power for effective photoporation and so as not to damage the targeted cells. A beam-shutter (Newport, UK, model 845HP-02) was used to provide the short exposure times (10 – 250 ms) which were required. The beam was then expanded with a lens relay telescope to fill the back of the microscope objective (×60; N.A. = 0.85). The focused beam radius, w0, was approximately 0.5 μm. A CCD camera was sited above the sample allowing direct observation and selection of individual cells for targeted transfection. Cells were positioned at the laser beam focus by manipulating an xyz translation stage on which the sample dish was placed. When focused onto the plasma membrane of a cell of interest, a small hole or pore was generated. These transient holes allowed femtolitre volumes of the surrounding medium, containing membrane impermeable substances such as the plasmid DNA, to enter the targeted cell.
3. Photoporation and transfection of CHO cells
For the cell transfection experiments, CHO cells were grown to sub-confluence in 30 mm diameter glass-bottomed culture dishes (usable area 23 mm; World Precision Instruments, Stevenage, UK) in 2 ml of Modified Eagles Medium (MEM) with 10% fetal calf serum (FCS) (Invitrogen, Paisley, UK), 20 units/ml of penicillin (Sigma, UK) and 20μg/ml of streptomycin (Sigma, UK) in a humidified atmosphere of 5% CO2/ 95% air at 37 °C. The cell monolayer was washed twice with OptiMEM (Invitrogen), and exposed to a 20 μl solution of OptiMEM containing 10 μg/ml pEGFPN2 plasmid (BD Biosciences, Oxford, UK), a plasmid encoding for green fluorescent protein (GFP). A 23 mm diameter type-1 coverslip (BDH, Poole UK) was then floated on top of this 20 μl solution and individual cells were irradiated with the laser from above. This configuration allowed cells to be irradiated on their non-adhered side, where they are exposed directly to the solution of plasmid DNA, while remaining in a semi-sterile environment (Fig. 2).
After treating the cells for between 10 to 250 ms at power levels between 50 to 225 mW at focus, the coverslip was removed, the monolayer gently washed twice in 90% MEM / 10% FCS, and the culture dish was returned to the incubator. 48 hours after treatment, photoporated cells were stained with 4’, 6-diamidino-2-phenylindole (DAPI - a blue nuclear dye that stains double stranded DNA) and viewed by fluorescent microscopy for GFP expression, allowing the transfection efficiency to be quantified. The transfection efficiency was calculated by dividing the number of cells expressing GFP at 48 hours by the total number of cells that were treated by the laser in a particular region of interest at 0 hours. Cells destroyed or irreversibly damaged as a result of the laser action were included in the data in order to provide a real and representative figure for the transfection efficiency of the femtosecond optical transfection technique. This appears to contradict some publications in the literature, where there has been a tendency to exclude such cells from transfection efficiency calculations.
48 hours after photoporation in the presence of plasmid DNA, fluorescent microscopy shows that transient GFP expression has been achieved and that cells expressing GFP are viable and display normal morphology, thereby confirming that optical transfection has occurred (Fig. 3). Spontaneous transfection (exposed to the GFP plasmid but not irradiated with the laser) can sometimes occur, but is negligibly low (<1 in 106 cells). In terms of controls, it was found that no GFP expression was observed when cells are exposed to the laser but not to the GFP plasmid, and that furthermore, no GFP expression was observed in cells exposed to either laser dosing or the GFP plasmid.
Before the full investigation into femtosecond optical transfection was carried out, the photoporation and transfection techniques described above were first repeated several times in a precursor study to provide knowledge and experience of the specific experimental requirements. It was therefore possible to develop a controlled and repeatable protocol for individual cell selection/treatment, high focusing accuracy, and the specific laser parameters corresponding to the desired cellular response. In particular, positioning of the highly-focused laser on the cell membrane requires sub-micron accuracy in the vertical axis, due to the related two-photon absorption process. This focusing accuracy was determined empirically during the precursor study by paying careful attention to the corresponding image plane of the targeted cells. It should be noted that, once laser parameters have been optimized for efficient photoporation, occasional cellular death as a result of laser action cannot be wholly avoided.
4. Observations during femtosecond photoporation
Successful photoporation of each targeted cell was determined by the visual response of the cell to the laser treatment. When successful, a small pore, or bubble, on the cell membrane was visible for no longer than the duration of the shutter window (A1-A3 in Fig. 4). If no pore was generated when the laser was exposed to the cell, it was assumed that photoporation, and therefore phototranslocation, had not occurred. This lack of cellular response was observed more often at lower power levels, and was employed to determine the threshold for successful photoporation, measured to be ~80 mW at focus with a 40 ms shutter time (corresponding to a localized fluence at the cell membrane of ~0.1 μJ/m2). The upper power limit of 225 mW at focus was set by the limits of the laser source and overall optical transmission of the microscope. The range of shutter times employed for this study were dictated by the lower limit of the shutter device (10 ms) and obvious cellular disruption or death (250 ms) even at very low laser power levels.
In photoporation it is possible to determine the likelihood of cell viability following laser treatment by observing the immediate cellular response to the laser action. Typically, cell exposure to shutter times greater than 100 ms resulted in a dramatic cell volume increase (up to 21% by visible area), the generation of an irreparable large hole (B1–B3 in Fig. 4), cell blebbing, granularity and death. Any dramatic size increase of a photoporated cell is likely to represent irreversible damage. Cell blebbing in the first few minutes after photoporation is an indication of cell stress and the onset of apoptosis.
The size of pore or bubble generated on the cell membrane is another visual key to determining cellular viability. Ideally, a pore approximately the size of the focused laser spot (diameter, d ~1 μm for a ×60, 1.25 N.A. microscope objective), together with its immediate re-closure following the closure of the shutter, provides the maximum chance for cell viability. In a study of 16 representative photoporated CHO cells, a range of hole sizes from ~0.8 μm2 (d ~1 μm) to ~75 μm2 (d ~10 μm) were generated. Cell morphology immediately following this procedure for each hole size was observed for a period of 10 minutes. It was observed that cells appeared healthy, with no signs of cell blebbing, granulation, or stress, when poration resulted in hole sizes between 0.8 μm2 and 3.9 μm2 (d ~2.2μm). For cells which were subject to larger bubble formation (28 μm2 to 75 μm2), cell viability was clearly compromised, as observed by morphological changes. By controlling laser fluence, exposure duration and focal location, similar regimes of operation have been also identified elsewhere in which cells could be selectively killed or kept viable immediately after bubble generation . In addition, bubbles with a lifetime of the order of a few seconds have been attributed to photochemical bond breaking as well as possible accumulative thermal effects . The appearance of such long-lived bubbles (i.e. B1-B3 in Fig. 4) has been reported as an indication of severe cell damage within the target region.
In order to illustrate and confirm that photoporation of cells by laser treatment acts to permeabilise the cell membrane, CHO cells were photoporated in the presence of 0.1% of the normally membrane impermeable dye, trypan blue. In Fig. 5, the cell indicated has been photoporated, and 30 seconds later the dye has penetrated the cell membrane through the laser-induced temporal pore. Phototranslocation of trypan blue has therefore occurred.
5. Femtosecond optical transfection efficiency
In order to determine a truly representative transfection efficiency for this relatively new technique, we performed the femtosecond optical transfection procedure (as described in section 3) on a large number of CHO cells in the presence of GFP plasmid DNA. In total, four thousand CHO cells were photoporated either in the presence of plasmid DNA, or in the absence of plasmid DNA as control experiments. CHO cells were transfected with a range of laser powers (from 50 mW to 225 mW) and exposed for a range of shutter times (from 10 ms to 250 ms). For each set of laser parameters, systematic control experiments were performed to exclude false results such as spontaneous transfection and contamination of either the cell line or the culture medium.
As discussed earlier, transfection efficiency, η, was calculated by dividing the number of cells expressing GFP at 48 hours by the total number of cells that are treated by the laser at 0 hours. In order to provide a real and representative figure for the transfection efficiency of the femtosecond optical transfection technique, cells destroyed or irreversibly damaged as a result of the laser action were included in the data. In Fig. 6, transfection efficiency is plotted as a function of fluence (the total deposited laser energy per unit area) at the cellular membrane, Φ. The fluence is calculated as the average laser power at focus multiplied by the exposure time, divided by the focal laser area (A = π, where w0 ~0.5 μm in this case). The main graph in Fig. 6 represents transfection efficiency, η, as a function of Φ, where Φ is expressed in units of μJ/cm2. For each value of Φ: the boxes represent the span of transfection efficiencies achieved; the data points represent the average transfection efficiency; the bars represent the standard error of the mean. In the inset in Fig. 6, each data point corresponds to one set of results, in which n = 100 cells were photoporated, n = 50 in the presence of plasmid DNA.
The transfection efficiency appears to depend strongly on the laser fluence employed for photoporation. Overall, the average transfection efficiency for cells treated with a fluence in the range of 0.5 – 2 μJ/cm2 was 36%. The average transfection efficiency at a specific fluence 1.2 μJ/cm2 was as high as 50 ± 10%. The maximum transfection efficiency achieved was around 80% at a fluence of 0.8 μJ/cm2. All data for transfection efficiency presented here so far includes cells killed or irreversibly damaged as a result of laser action. This appears to contradict some publications in the literature, where there has been a tendency to exclude non-viable cells in order to quote ‘corrected’ transfection efficiencies.
6. Viability of photoporated cells
To assess the extent to which transfection efficiency is compromised due to the loss of cells resulting from the combination of natural processes and laser treatment, the viability of photoporated cells was quantified 2 hours after photoporation. A highly specified region of interest (~100 × 1000μm) was first micromachined on a 25 mm diameter polystyrene culture dish by lowering the focus of the femtosecond laser and ablating lines on the polystyrene surface. The surface was then washed three times in 90% MEM / 10% FCS, and CHO cells were seeded and grown to confluence 24 hours later on this surface. All cells within the region of interest (n ~200) were then photoporated at fluences between 1 – 2 μJ/cm2 (the optimum laser fluence for high transfection efficiency) and viability assessed 2 hours later by the ability of cells to exclude 0.1% of the membrane impermeable dye, trypan blue in phosphate buffered saline (pH = 7.4) (PBS; Invitrogen). An adjacent marked region of equal size was not treated by the laser, but instead used to determine natural cellular viability.
Cells appearing to have taken up the blue dye after 2 hours were considered non-viable, as their membrane integrity had been terminally compromised. This procedure was repeated three times for a viability study involving the photoporation of 600 cells in total. This allowed the viability of cells treated to a particular laser dose to be corrected for the background loss of viability in a normal cell monolayer. Any cells observed to have detached from the culture dish during the 2 hour period were considered non-viable and included in the data.
The overall cellular viability (the number of live cells divided by total number of cells) for a region of cells being treated by the laser was found to be 65 ± 7%. In the non-laser control [the lower region in Fig. 7(b)], the natural cellular viability was found to be 92 ± 3%. Combining these figures provides an overall corrected cellular viability of Vc = 70 ± 8% for CHO cells undergoing femtosecond photoporation. Vc may then be applied as a correction factor to the transfection efficiency, η, to provide the corrected transfection efficiency, ηc = (100/Vc)η. This corrected transfection efficiency, ηc, for the range of fluences between 0.5 – 2 μJ/cm2 is then 51%. At a fluence of 1.2 μJ/cm2, ηc rises to 71%.
7. Discussion and conclusions
The underlying cytophysiological and related mechanisms by which the a pulsed laser creates a hole in the cell membrane have been attributed to numerous similar effects, such as photodisruption , dissection , disassociation of water molecules leading to the presence of reactive oxygen species , and to localized plasmas resulting from laser-induced optical breakdown (LIOB) . In our observations, the action of photoporation is certainly dominated by a nonlinear process and depends strongly on laser intensity. The threshold intensity required for optical breakdown in water is ~1013 W/cm2  and there is some agreement in the literature that, assuming cellular membranes are rich in water and for lasers with μJ pulse energies, LIOB is the dominant mechanism. Such threshold intensities can be reached with femtosecond lasers with nJ pulse energies, but in these cases it can not be said with any certainty whether or not LIOB is responsible. A mechanism has recently been proposed for femtosecond pulse trains at MHz-repetition-rates with energies below the threshold for LIOB. Vogel and co-workers  describe intracellular ablation by the presence of low-density plasmas which rely on cumulative free-electron-mediated chemical effects. Specifically, transient membrane permeabilisation for the purposes of gene transfection requires a much larger laser dose than chromosome dissection, and therefore higher degree of photochemical bond-breaking. The corresponding laser parameters for such photoporation are still within the regime of free-electron-mediated chemical effects, but may also involve contributions from cumulative heating effects.
In this paper we have reported the first detailed study into transfection efficiency and cell viability for the technique of femtosecond optical transfection. Photoporation of four thousand Chinese Hamster ovary cells at a range of laser fluences was undertaken in the largest optical transfection study of individual cells reported to date. Overall, we determine an average femtosecond transfection efficiency for cells treated with a fluence in the range of 0.5 – 2 μJ/cm2 of 36 ± 5%. The average transfection efficiency at a specific fluence of 1.2 μJ/cm2 was found to be 50 ± 10%.
Cellular viability as a result of laser treatment was found to be 65 ± 7%. Natural cellular viability was found to be 92 ± 3%, providing a corrected cellular viability 70 ± 8% for CHO cells. In order to present our data in a form consistent with the literature, we applied the observed cellular viability to obtain corrected transfection efficiencies, ηc. For the range of fluences between 0.5 – 2 μJ/cm2, ηc was calculated to be 51 ± 9% and at the specific fluence of 1.2 μJ/cm2, ηc was found to be 71 ± 16%.
Even taking account of cell losses as a result of the laser treatment, we find that optical transfection, although an exciting technique, is not 100% as alluded to in previous work  - which is also true for any other transfection technique. Although the applied laser fluence can be selected in order to maximize the chances of optical transfection occurring, individual cellular viability of the targeted cell cannot be guaranteed. In addition, the transfection efficiency of the many cells that survive the photoporation process is compromised by the requirements of the treated cell to then undergo the nuclear translocation, transcription, and translation processes. As such, we consider a transfection efficiency that takes account of cellular viability of around 50% to be a true representation of the usefulness of this technique.
D.S., B.A., F.G., and K.D. contributed equally to the work presented. We thank the UK Engineering and Physical Sciences Research Council and the Medical Research Council for funding, and we are grateful to L. Paterson and M. Comrie for their contributions to the transfection experiments. The assistance of A. D. McRobbie and N. K. Metzger with the titanium sapphire laser has also been greatly appreciated.
References and links
1. S. Mehier-Humbert and R. H. Guy, “Physical methods for gene transfer: Improving the kinetics of gene delivery into cells,” Adv. Drug Delivery Rev. 57, 733–753 (2005). [CrossRef]
3. G. Palumbo, M. Caruso, E. Crescenzi, M. F. Tecce, G. Roberti, and A. Colasanti, “Targeted gene transfer in eucaryotic cells by dye-assisted laser optoporation,” J. Photochem. Photobiol. B 36, 41–46 (1996). [CrossRef] [PubMed]
4. L. Paterson, B. Agate, M. Comrie, R. Ferguson, T. K. Lake, J. E. Morris, A. E. Carruthers, C. T. A. Brown, W. Sibbett, P. E. Bryant, F. Gunn-Moore, A. C. Riches, and K. Dholakia, “Photoporation and cell transfection using a violet diode laser,” Opt. Express 13, 595–600 (2005). [CrossRef] [PubMed]
5. M. Tsukakoshi, S. Kurata, Y. Nomiya, Y. Ikawa, and T. Kasuya, “A novel method of DNA transfection by laser microbeam cell surgery,” Appl. Phys. B 35, 135–140 (1984). [CrossRef]
6. Y. Shirahata, N. Ohkohchi, H. Itagak, and S. Satomi, “New technique for gene transfection using laser irradiation,” J. Inv. Med. 49, 184–190 (2001). [CrossRef]
7. J. S. Soughayer, T. Krasieva, S. C. Jacobson, J. M. Ramsey, B. J. Tromberg, and N. L. Allbritton, “Characterization of cellular optoporation with distance,” Anal. Chem. 72, 1342–1347 (2000). [CrossRef] [PubMed]
8. S. Sagi, T. Knoll, L. Trojan, A. Schaaf, P. Alken, and M. S. Michel, “Gene delivery into prostate cancer cells by holmium laser application,” Prostate Cancer Prostatic Dis. 6, 127–130 (2003). [CrossRef] [PubMed]
9. S. K. Mohanty, M. Sharma, and P. K. Gupta, “Laser-assisted microinjection into targeted animal cells,” Biotech. Lett. 25, 895–899 (2003). [CrossRef]
11. U. K. Tirlapur and K. Konig, “Femtosecond near-infrared laser pulses as a versatile non- invasive tool for intra-tissue nanoprocessing in plants without compromising viability,” Plant J. 31, 365–374 (2002). [CrossRef] [PubMed]
12. E. Zeira, A. Manevitch, A. Khatchatouriants, O. Pappo, E. Hyam, M. Darash-Yahana, E. Tavor, A. Honigman, A. Lewis, and E. Galun, “Femtosecond infrared laser - an efficient and safe in vivo gene delivery system for prolonged expression,” Mol. Therapy 8, 342–350 (2003). [CrossRef]
13. A. Vogel, J. Noack, G. Huttmann, and G. Paultauf, “Mechanisms of femtosecond laser nanosurgery of cells and tissues,” Appl. Phys. B 81, 1015–1047 (2005). [CrossRef]
14. D. Stevenson, B. Agate, L. Paterson, T. K. Lake, M. Comrie, C. T. A. Brown, A. C. Riches, P. E. Bryant, W. Sibbett, F. Gunn-Moore, and K. Dholakia, “Optical transfection of mammalian cells,” presented at SPIE Photonics Europe, Strasbourg, France, 3–7 April 2006.
15. M. J. Zohdy, C. Tse, J. Y. Ye, and M. O’Donnell, “Optical and acoustic detection of laser-generated microbubbles in single cells,” IEEE Trans. Ultrason. Ferroelectr. Freq. Control 53, 117–125 (2006). [CrossRef] [PubMed]
16. A. Heisterkamp and H. Lubatschowski, “Subcellular photodisruption,”Femtosecond technology for technical and medical applications, Top. Appl. Phys. 96, 227–232 (2004). [CrossRef]
17. A. Heisterkamp, I. Z. Maxwell, E. Mazur, J. M. Underwood, J. A. Nickerson, S. Kumar, and D. E. Ingber, “Pulse energy dependence of subcellular dissection by femtosecond laser pulses,” Opt. Express 13, 3690–3696 (2005). [CrossRef] [PubMed]
18. U. K. Tirlapur, K. Konig, C. Peuckert, R. Krieg, and K. J. Halbhuber, “Femtosecond near-infrared laser pulses elicit generation of reactive oxygen species in mammalian cells leading to apoptosis-like death,” Exp. Cell Res. 263, 88–97 (2001). [CrossRef] [PubMed]