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Abstract
The confocal fluorescence microscope is an essential live cell imaging tool in bioscience research. Several experimental investigations in the field of biomedical research require a dedicated confocal fluorescence microscope. However, commercial confocal microscopes are prohibitively expensive for many individual laboratories and they often have an inflexible design not amenable to user desired modifications. Here we report on the design, development, and calibration of a cost-effective dual channel confocal fluorescence microscope that can capture two biological events simultaneously. The microscope is successfully employed to image and study the simultaneously occurring active and passive transport of molecules across the nuclear membrane. Passive diffusion of FITC labelled dextran molecules are monitored along with the active transport of gold nanoparticles of diameter 20 nm in the time-lapse imaging mode. The experiments carried out in digitonin permeabilized HeLa cells indicate that both active and passive nuclear transport pathways coexist together.
© 2021 Optical Society of America under the terms of the OSA Open Access Publishing Agreement
1. Introduction
Confocal fluorescence microscopy is one of the most important developments in the field of optical microscopy. Three-dimensional imaging in confocal microscopy is achieved by capturing thin cross-sectional images across the specimen by optical sectioning and combining them [1]. The technique uses point illumination by a laser source and makes use of spatial filtering to collect fluorescence signals from the focal plane of the specimen, thereby providing better contrast and resolution [2]. The laser scanning method employed in modern confocal microscopes to raster scan the beam over the sample enables fast image acquisition and allows monitoring of fast biological processes. The improved axial resolution and the ability to nondestructively capture three-dimensional images of live specimens make confocal fluorescence microscopy the favorite imaging tool in biomedical research.
Introduced in 1957 by Marvin Minsky, the concept of confocal imaging is widely used in various areas of biology [3], biomedicine [4,5], food science [6], and material sciences [7]. The availability of a variety of biocompatible fluorochromes in a wide range of spectral regions has provided a boost to the application of confocal microscope in live cell imaging. Fluorescence spectroscopic techniques such as fluorescence resonance energy transfer (FRET) [8,9] fluorescence lifetime imaging (FLIM) [10] and fluorescence recovery after photobleaching (FRAP) [11] has further broadened the application of confocal microscope in biomedical research. Though many new optical microscopic techniques catering to specific applications have been developed in recent years [12–14], confocal fluorescence microscopy remains the most favored imaging tool due to its simplicity and wide range of applications.
Confocal fluorescence microscopy became widely popular in biomedical research after the introduction of the first commercial microscope in 1987 by W. B. Amos, J. G. White, and M. Fordham [15–17]. However, the high purchase cost of a commercial confocal microscope precludes its accessibility to many research laboratories. The design of commercial confocal microscopes cannot be easily modified by the user which is often a dampener. Many laboratories require dedicated confocal microscopes for longtime imaging and monitoring of biological systems and may want to configure it as per specific laboratory requirements. In this context, an in-house constructed confocal laser scanning microscope with a small budget and without compromising on the imaging capability is an attractive alternative [18–21]. We have recently constructed a cost-effective single-channel confocal laser scanning microscope (CLSM) with the help of laser and optical components available in the laboratory [21]. The microscope which can capture a single biological event is also used for multiphoton [14] and photothermal microscopy applications [13] with minimal modifications. Here we report on the development of a dual-channel confocal fluorescence microscope with enhanced imaging capability and capable of simultaneously monitoring two biological events. Additional channels to the microscope can be easily added depending on the user requirement. One of the challenges faced by many in the construction of a CLSM is the development of operating software to interface hardware components and for data acquisition. In this work we make use of an open-source software, ‘ScanImage’ to control the CLSM as well as for image acquisition [22]. The image analysis is done with the help of an open-source image processing program ‘ImageJ’ The in-house constructed confocal microscope is successfully employed to study biomolecular transport through digitonin permeabilized HeLa cell nuclear membranes [23]. In particular, we make use of time-lapse confocal fluorescence microscopy to image the passive and active nuclear import of two model nanoparticles, FITC-dextran and gold nanoparticles respectively, through the nuclear pore complexes [24–28].
2. Methods
2.1. Image formation in confocal microscopy
The method of image formation in a confocal laser scanning microscope is entirely different from that of a traditional wide-field optical microscope. In the case of the wide-field microscope, the whole specimen is illuminated with a broadband light source and the signal is collected using a flat 2D detector such as a CCD camera or directly viewed by the eye. In this scheme of illumination and detection, scattered light and out-of-focus light are detected along with the desired signal from the focal plane. This severely affects the resolution of the microscope and its ability to capture fine details of the specimen. In confocal microscopes, point-by-point illuminations is done by tightly focusing the excitation beam to a diffraction-limited spot in the specimen kept at the objective focal plane. In CLSM, the required area of the specimen is imaged by moving the focal spot in the x-y plane using a scan mirror system. A scan lens kept after the scan mirrors along with the tube lens of the microscope provide a collimated input beam to the objective. The point detection is done by making use of a pinhole aperture in front of the detector, at the conjugate focal plane of the objective lens. A detector lens is used to focus the fluorescence light to the aperture. The aperture essentially blocks out-of-focus light and allows only the light from the focal plane to reach the detector. Several major technological developments like the introduction of multiple wavelength lasers, highly sensitive photodetectors and faster data acquisition and processing systems have helped to significantly improve the performance of current confocal microscopes when compared to Minsky’s original design.
There is extensive discussion on the theoretical resolution of confocal fluorescence microscopes [1,29–31] and the relation between the resolution, the excitation wavelength, and the numerical aperture. The more appropriate and straightforward diffraction-limited lateral resolution (rlateral) of a CLSM is
where ${\lambda _{exc}}$ is the excitation wavelength of the laser used and NA is the numerical aperture of the objective used [29].Experimentally the resolution of the confocal microscope is determined by measuring the full width at half maximum (FWHM) of the lateral point-spread function. The point spread function of CLSM is the three-dimensional diffraction pattern of a point-like object under the microscope. Typically, a fluorescent nanoparticle having a size much less than the diffraction limit can be used to study the point spread function. The diffraction pattern by a point-like object is also known as the Airy pattern and the central bright ring is called Airy disc. The diameter of the projected Airy disc at the confocal aperture plane of CLSM can be calculated by the equation [20,21]
where, ${d_p}$ is the diameter of the Airy disc, ${f_D}\; $ is the focal length of the detector lens, ${f_{SL}}$ is the focal length of the scan lens, ${M_{obj}}$ is the magnification of the objective lens. In the present experiment, an objective of magnification 60X with NA of 1.25 is used along with a scan lens of focal length 25 mm. The focal length of the detector lens used is 15 cm for the 488 nm excitation and 25 cm for the 543 nm excitation. Thus the airy disc diameter for 488 nm excitation is 171 µm and that for 543 nm excitation is 318 µm.The axial resolution of CLSM is given by the relation [29]
where, n is the refractive index of the medium. Here we have used an oil immersion objective and the refractive index of the immersion oil used is 1.518. Thus, the theoretical axial resolution for the 488 nm excitation is 654 nm. The corresponding lateral resolution given by Eq. (1) is 200 nm.2.2. Design and construction of the confocal microscope:
2.2.1. Hardware for microscope construction
The components used in the construction of the confocal laser scanning microscope are listed in Table 1 along with an approximate cost. In brief, we have used an inverted optical microscope (Olympus IX 71) with a 60X oil immersion objective (NA = 1.25) (UPlanFLN 60 X, Olympus) for setting up of the CLSM. One of the considerations to keep in mind while choosing the microscope is the number of input and output ports required for any future upgradation to different configurations such as a multiphoton microscope. The fluorescently labelled specimen under study is kept at the focus of the microscope objective. It is excited by the laser entering through the left side port of the microscope and fluorescence emitted by the specimen is collected through the same port. Two lasers, an Ar+ laser having wavelength 488 nm and a He-Ne laser having wavelength 543 nm, are used as the sources for the two excitation channels. One may choose low-cost diode lasers readily available at different wavelengths to construct a compact confocal microscope. A galvanometric mirror scanner (GVS002/M, Thorlabs) is used to raster scan the beam over the sample. GVS002/M is a dual-axis galvo system that contains two mutually orthogonal X and Y silver-coated scan mirrors. The X mirror moves the beam along X-axis and the slow-moving Y mirror moves the beam along a perpendicular direction to complete a raster pattern of the scan. The movement pattern of the scan mirror depends upon the analog signal given by the controller. In our experiment, a bidirectional scanning method is employed to scan the required area of the specimen. In bidirectional scanning, the beam sweeps from left to right and returns to the next line and data is collected in both directions of motion of the scan mirror. A triangular voltage waveform is applied to the X scan-mirror and a sawtooth waveform is applied to Y scan-mirror. The frequency of the wave form applied to the X scan-mirror is 256 times that of the Yscan-mirror. Thus to acquire a 512 x 512 pixel image both mirrors are simultaneously moving in such a way that the X mirror completes 256 cycles of movement during one cycle of movement of the Y mirror. The two distinct fluorescence signals from the two different fluorophores retract the excitation path and are filtered through dichroic mirrors and filters and finally detected by two different photomultiplier tubes (PMT) (H7732-10, Hamamatsu). The fluorescence signal from the sample is coupled to the PMT via optical fibers which act as the confocal aperture for spatial filtering. A transimpedance amplifier (C12419, Hamamatsu) is used to convert PMT output current to voltage. A data acquisition card (NI USB 6356 DAQ) is used for providing the necessary voltage for the scan mirrors and it simultaneously acquires data from the PMT’s. The PMT output is recorded in synchronization with the mirror position (and thus the beam position on the sample) with the help of an imaging and data acquisition software. An open-source program based on Matlab software, ‘ScanImage’ (Vidrio technologies, version 5.1) is used for controlling the scan mirrors and acquiring the image. The software is compatible with Windows 10 (64-bit) operating system, 64- bit Matlab 2015 version, and the NI Driver DAQmx 15.1.
Table 1. List of components used for the construction of dual channel CLSM
2.2.2. Laser beam alignment
A detailed schematic of the dual channel laser scanning confocal microscope is given in Fig. 1. The laser lines selected for fluorescence excitation are 488 nm (channel 1, Ar+ laser) and 543 nm (channel 2, He-Ne laser). Both laser beams are expanded and collimated using a telescopic arrangement (T) as shown in Fig. 1. This beam expander ensures that a collimated beam of size less than 5 mm (size of scan mirror) is reflected off the scan mirror. Both laser beams are made collinear with the help of a dichroic mirror, DM1 (DMLP505, Thorlabs). DM1 allows wavelengths above 505 nm to pass through and reflects those below 505 nm. Mirrors M2, M3, and dichroic mirror DM2 (ZT488/543rpc, Chroma) are used to direct the laser beams to the scan mirrors. Dichroic mirror DM2 is a multi-band dichroic beam splitter that reflects 488 nm and 543 nm laser lines and directs it to the scan mirror. At the same time, DM2 allows the fluorescence signal having a wavelength between 490 nm to 540 nm as well as a wavelength above 550 nm to pass through it. The reflected light from the scan mirrors is directed to the microscope's left-side port through a scan lens, SL (UIS2-WHN10X/22, Olympus). The focal length of the scan lens is 25 mm and is kept at a distance of 127 mm from the microscope's left-side port so that the front focal plane of the scan lens coincides with the primary image plane. The scan mirrors and scan lens are mounted on an XYZ micrometer translation stage to facilitate the alignment.
Fig. 1. Optical layout of dual fluorescence channel confocal microscopy. DM1,2,3: Dichroic Mirrors BPF1,2,3: Bandpass filters, S: Shutter, T: Telescope, M 1,2,3,4: Mirrors, SL: Scan lens, XS, YS: X and Y scan mirrors, BS: Beam splitter, PMT: Photomultiplier Tube, CA1,2: Confocal Apertures, DL1,2: Detection lenses, DAQ: Data Acquisition Card, Ar+ 488: Ar+ laser having an emission wavelength of 488 nm, He-Ne 543: He-Ne laser having an emission wavelength of 543 nm. Additional excitation lines can be added in parallel to these laser lines effortlessly.
The optics of the microscope directs the laser beam, expanded to a size of 17 mm, to the back aperture of the objective which is 10 mm in diameter. This overfilling is done for assuring a uniform beam entry to the objective while scanning. The laser beam reaches the specimen kept at the focal plane of the objective and the emitted fluorescence returns in the same path and is de-scanned by the scan mirrors before passing through the dichroic mirror DM2. The dual-band dichroic mirror DM2 passes the fluorescence emission from the two excitations while blocking the excitation lasers. The emitted fluorescence is further filtered through a dual bandpass filter BPF1 (ZET 405/488/543 nm, Chroma) and reaches the dichroic mirror DM3 (FF 553-SDi01-25, Semrock) which transmits the fluorescence from 488 nm excitation and reflects the fluorescence emitted from the 543 nm excitation. Mirror M4 is used to direct the fluorescence signal for 488 nm, transmitted through DM3, towards the detection lens DL1 (focal length 15 cm). The bandpass filter BPF3 (ET 520/40 m, Chroma) is used in the detection path for further filtering the fluorescence signal. The detection lens DL1 focuses the fluorescence signal to the optical fiber, CA1 (diameter = 200 µm M72L01, Thorlabs) connected to the 488 nm PMT. The fluorescence signal from 543 nm excitation is further filtered by using a bandpass filter BPF2 (FELH0550, Thorlabs) and directed towards the detection lens DL2 (focal length=25 cm). The detection lens focuses the fluorescence signal to the optical fiber, CA2 (diameter = 400 µm) connected to the 543 nm PMT. The optical fiber transmits only the light falling on its core and couples it to the detector. Thus, the optical fiber act as the confocal aperture which removes the out-of-focus signal [20]. Here the core diameter of the fiber is the diameter of the confocal aperture. The amplified analog signal from PMT is converted to a digital signal by a data acquisition card and is converted into an image using the ScanImage program.
Figure 2 further illustrates the optical layout of the scanning system along with the beam path from the scan mirrors to the microscope. The beam paths corresponding to two nearby points on the specimen plane are indicated by the blue and red lines. The figure depicts the specimen plane and its primary image plane which lies between the tube lens and scan lens, back focal plane of the objective (which coincides with the rear aperture of the objective) and its conjugate focal plane. The X and Y scan mirrors are closely spaced (close-coupled) and is placed in such a way that the back focal plane of the scan lens is at the center of the X and Y scan mirrors. The back focal plane of the scan lens is a plane conjugate with the back focal plane of the objective. Pivoting the beam at a stationary point on the conjugate plane minimizes the movement of the laser beam at the rear aperture of the objective as illustrated in the figure. This will prevent illumination fall-offs at the rear aperture of the objective during the scan cycle [32]. It may be noted that all light beams intersect at the rear aperture of the objective (and the conjugate focal plane) at an angle that is a function of the position of the source point in the specimen plane. The positioning of the scan mirror at the conjugate focal plane with its pivot point on the optic axis produces a linear motion of the focused beam spot on the specimen. The fluorescence signal from the sample retraces back along the excitation path and is descanned by the scan mirrors as shown in the figure.
Fig. 2. Optical layout of the scanning system along with the beam path from the scan mirrors to the microscope. The blue and red colored lines indicate the beam paths corresponding to two nearby points on the specimen plane.
2.3. Nuclear transport protocol
We have chosen the HeLa cell line, a cervical cancer cell line, for the study of the transport of dye labelled molecules across the nuclear pore complexes (NPC). The molecule chosen for the passive nuclear transport were Fluorescein isothiocyanate (FITC) dye labelled Dextran molecules having a molecular weight of 10 KDa (Model: FD10S, Sigma Aldrich Chemicals Pvt. Ltd). The imaging for the passive nuclear transport is done using channel 1 of the CLSM. CY3 (Cyanine) dye-labeled Nuclear Localization Signal (NLS) tagged gold nanoparticles (GNPs) (CF11-20-CY3-NLS-DIH-50, Nanopartz Inc) having a mean diameter of 20 nm is used for the active nuclear transport study. Channel 2 of the dual-channel CLSM is used for active nuclear transport imaging.
2.3.1. Cell culture
We have purchased the HeLa cell line for the current study from the Cell Repository division of the National Centre for Cell Science (NCCS), Pune, India. The cells are cultured in a complete cell culture medium in which 90% Dulbecco’s Modified Eagles medium (DMEM) is supplemented with 10% Fetal Bovine Serum (FBS). The cultures are maintained in a humidified incubator at 37°C temperature and 5% CO2 atmosphere. The cells for the transport studies are grown in imaging chambers. These imaging chambers are prepared by drilling a 6 mm hole at the bottom of Petri dishes (35 × 15 mm, 460035, TARSONS) and fixing a coverslip thereby melting parafilm using a Bunsen burner.
2.3.2. Cell permeabilization and Nuclear import Assay
2.3.2.1. Solutions and reagents:
The buffer solution for the nuclear transport called transport buffer is prepared from 20 mM HEPES, PH 7.3, 110 mM KAc, 5 mM NaAc, 2 mM MgAc, 0.5 mM EGTA, 2 mM DTT, 1 μg/ml of aprotinin, leupeptin, and pepstatin (protease inhibitors). The optimized concentration of 40 µg/ml digitonin in transport buffer is used for cell permeabilization. The digitonin permeabilization is carried out for the optimized time period of 5 minutes. Rabbit reticulate lysate (RRL, L4960, Promega) is used as the artificial cytoplasm to fulfill the requirement of cytosolic factors necessary for nuclear transport [33]. The RRL has to be dialyzed against the transport buffer before using it in the nuclear transport studies. 50% of the final solution for the nuclear transport studies consists of RRL. The remaining 50% contains the desired concentration of import cargo molecule prepared in transport buffer. For active nuclear transport studies, the import mixture is supplemented with an additional energy regenerating system (1 mM Mg ATP, 1 mM GTP, 5 mM creatine phosphate, 20 U/ml creatine phosphokinase). In the present study, the final import mixture for the simultaneous active and passive nuclear transport contains 0.5 mg/ml of 10 KDa FD molecules, 0.5 mg/ml of 20 nm gold nanoparticle tagged with NLS and labelled with CY3 dye, energy regenerating system, and remaining 50% volume is RRL. The detailed protocol for nuclear transport study is given as
- 1. Seed around 20000 cells in confocal dishes 18 to 24 hrs. prior to the experiment.
- 2. On the day of the experiment replace the culture medium with a fresh cell culture medium around 2 hrs. prior to the experiment.
- 3. Prepare all solution mixtures for the experiment in enough volume freshly on the day of the experiment.
- 4. Remove cell culture medium from the confocal dish and wash two times with transport buffer.
- 5. Permeabilize the cells by incubating with 40 µg/ml of digitonin solution for 5 minutes at room temperature.
- 6. Stop permeabilization process by aspirating digitonin and replacing it with complete transport buffer. Wash the cells three times with transport buffer.
- 7. Keep imaging chambers with cells in dual-channel CLSM and focus on the equatorial plane of the nucleus.
- 8. Replace transport buffer with import mixture and start imaging in the time-lapse mode of the confocal microscope.
3. Results and discussions
3.1. Calibration of the microscope
The size of the scan area for a given galvanometer voltage is determined by imaging the 1951 USAF resolution test target (Thorlabs, R3L3S1N - Negative). The test target contains a series of horizontal and vertical lines arranged in 10 groups of varying line spacing and are used to calibrate the imaging system. Figure 3. shows the group 7- element 6 (smallest pair of vertical and horizontal bars) captured by the dual-channel confocal laser scanning microscope. A 512 x 512 pixel image is taken by applying a voltage of 5 V to the scanner. The red line marked across a line pair (the black and a white bar) of the 6th element denote the specified length of 4.38 µm and have a measured pixel length of 21.7 pixels. Based on this we have calibrated our area of scanning for a voltage of 5 V to 103 µm X 103 µm.
Fig. 3. Calibration of CLSM by 1951 USAF Resolution test target. The image is obtained by providing 5 V to the scan mirrors. The smallest pair of horizontal and vertical bars corresponding to the 6th element of group 7. The length of the red line in the figure denotes the specified length of a line pair (black and a white bar) and is 4.38 µm corresponding to 21.7 pixels. The total image is 512 X 512 pixels corresponding to the imaging area of 103 × 103 µm.
The lateral and axial resolution of the microscope is determined by imaging dye-labeled sub-resolution polystyrene beads. Dye labeled polystyrene beads (Model: L9904, Sigma-Aldrich Chemicals Pvt. Ltd., USA) of 100 nm size are used for determining the resolution of the microscope. The stock concentration was 2.5% solids in 1 ml of solution. The stock solution is further diluted 106 times in double-distilled water and seeded in the imaging chamber. Figure 4. (a) shows the confocal images of 100 nm polybeads acquired using 488 nm line of the Ar: ion laser as the excitation source. The 512 × 512 pixels image of the scan area of 10.5 µm x 10.5 µm is acquired in 20 seconds. The lateral resolution of the microscope is measured from the point spread function (PSF) obtained from the image of a single 100 nm polystyrene bead shown in Fig. 3 (a). The line intensity profile through the midpoint of a polybead image is shown in Fig. 4 (b). The full width at half maximum of the profile is corresponding to a lateral resolution of 230 ± 30 nm in agreement with the predicted theoretical resolution. Here we may note that the experimentally measured PSF can depend upon on the size of the bead used and on different instrumental variables of the microscope [34]. Ideally the image of a point like object should be used to estimate the PSF. However smaller size beads will give rise to larger noise and photobleaching and would be experimentally challenging to determine the PSF by imaging a point-like bead. As a compromise one may choose a bead having a size much less than the expected resolution of the microscope, but large enough to provide good signal to noise ratio in imaging. PSF can also be affected by the optical alignment and the aberrations in the optical components used. Thus it is always good to use the experimentally determined PSF to reconstruct the image rather than the theoretical PSF.
Fig. 4. (a) The CLSM image (512 X 512 pixel) of a 100 nm dye-labeled polystyrene beads excited with 488 nm (b) The lateral and (c) axial line intensity profile of 100 nm polystyrene beads
The axial resolution of the microscope is determined from the axial intensity profile of 100 nm polystyrene beads obtained from multiple cross-sectional images in the axial direction. The axial movement of the objective in the order of nanometer-scale is done with the help of a piezoelectric objective mount (PFM 450E, Thorlabs) with a paired controller (PPC001, Thorlabs). The movement of the Piezo scanner is controlled by Kinesis software (Thorlabs). Figure 4(c) shows the axial intensity profile obtained by averaging over 10 polystyrene beads. The axial resolution of the microscope determined from the full width at half maximum of the profile is 800 ± 40 nm . Here we may note that the experimental axial resolution depends critically on the core diameter optical fiber used to couple the fluorescence to the detector. For the best resolution one should use an optical fiber having a core diameter equal to the diameter of the airy disc. As discussed in section 2.1, the estimated airy disc diameter for the 488 nm channel of our microscope is 171 µm. In our experiments we have used an optical fiber readily available in the lab having a core diameter of 200 µm and the experimentally determined axial resolution is less than the theoretical value of 654 nm.
Standard biological test slides are used for the verification of microscope performance. The microscope’s dual-channel performance is verified by imaging a multiple stained test slide containing Muntjac skin fibroblast cells (Model No. F-36925, Invitrogen Corporation, USA). The actin filaments in fluocells are stained with Alexa Fluor 488 phalloidin. Mitochondria are labeled with an anti–OxPhos Complex V inhibitor protein mouse monoclonal antibody in conjunction with Alexa Fluor 555 goat anti-mouse IgG. The confocal fluorescence images of the sample are acquired by simultaneously exciting the sample with the two laser wavelengths, 488 nm, and 543 nm. Images of the actin filaments in the cell acquired using the fluorescence emission from the 488 nm excitation are displayed in one of the channels. Corresponding images of mitochondria simultaneously acquired using the 543 nm excitation are displayed in the second channel of the confocal microscope. Figure 5. (a) Shows confocal image of actin filaments and Fig. 5. (b) shows confocal image of mitochondria of Muntjac skin fibroblast cells. The 512 x 512 pixel images of the scan area of 103 µm × 103 µm are acquired in 20 seconds, with a pixel dwell time of 76 µs. Figure 5. (c) is a reconstructed image obtained by merging the above two images. The signal to noise ratio of the acquired image is calculated as the ratio of maximum intensity of the image signal to the standard deviation of the background is 770.
Fig. 5. The simultaneous dual-channel imaging capability of CLSM by imaging fluorescently labeled Muntjac skin fibroblast cells (a) Images of Alexa Fluor 488 phalloidin labelled actin filaments excited with 488 nm laser beam (b) Alexa Fluor 555 labelled Mitochondria excited with 543 nm laser beam(c) The combined dual-channel confocal image of Muntjac skin fibroblast cells.
The optical sectioning capability of the microscope is verified by imaging multiple dye-labeled pollen grains (Model No. 304264, Carolina Biologicals, USA). In the dual-channel mode, the pollen grains are excited by two different wavelengths (488 nm and 543 nm) and the fluorescence images are acquired simultaneously and displayed in two separate channel displays. Optical sections of the pollen grain are taken by moving the objective in 1 µm steps along the z-axis. In Fig. 6 we show 12 cross-sections out of 36 cross-sectional images of the pollen grain acquired in channel 2. Figure 7 shows the 3D image of the pollen grain reconstructed from cross-sectional images using ImageJ, which is open-source software. The cross sectional images of the pollen grain acquired using 488 nm channel (not shown) also looks similar in size and shape to that acquired by the 543 nm channel.
Fig. 6. Cross-sectional CLSM images taken through a dye-labeled pollen grain excited with 543 nm laser. Each cross-sectional image represents the different focal planes across the pollen grain by an approximate distance of 3 μm.
3.2. Live cell imaging and nuclear import studies
The nuclear pore complexes in the nuclear membrane are the sole gateway of translocations between the cytoplasm and the nucleus [35–37]. The functioning of the nuclear pore complexes is quite complex, nuclear pore complexes allow the passage of small molecules via passive diffusion while blocking or allowing larger molecules inside depending on whether or not they carry the nuclear localization signal [25]. It is amazing to note that each nuclear pore complex in the nucleus is able to handle around 1000 molecular translocation events per second [38]. It would be interesting to see the two transport pathways, the active and passive nuclear transport pathways taking place simultaneously through the same nuclear pore complexes. In the present experiment, we make use of the in-house constructed confocal microscope to image the passive as well as the active transport of certain model biomolecules taking place at the same time through the nuclear membrane. FITC labeled dextran molecules having a molecular weight of 10 KDa are used to study passive nuclear transport. The dextran molecules are inert and have an approximate size of 4.6 nm diameter, below the reported passive permeability limit of 10 nm [24]. Gold nanoparticles having 20 nm diameter labelled with CY3 dye and nuclear localization signal (GNP-CY3-NLS) are used to study the active nuclear transport. This size of the gold nanoparticles is well below the size limit for active nuclear transport which is around 30 nm, but larger than the passive limit. The experiments are carried out on digitonin permeabilized HeLa cell nuclei in the time-lapse confocal imaging scheme. In order to image the passive and active nuclear transport simultaneously, an import mixture containing 0.5 mg/ml of FITC dextran and 0.5 mg/ml of GNP-CY3-NLS and RRL with additional energy regenerating systems is added to the imaging chamber in which the permeabilized HeLa cell nuclei are maintained. The exogenous cytosol RRL in the import mixture mimic as an artificial cytoplasm and provides all cytosolic transport factors necessary for the nuclear transport. The nuclear import of the transport cargo, which includes both FITC-dextran and GNP-CY3-NLS, is monitored in the two channels of the confocal fluorescence microscope. Images of the central cross-section of the nucleus is captured every 2.2 seconds after the addition of the import mixture. The imaging chamber in each experiment typically contains 6-7 intact nuclei in the scanning area and the experiment is repeated on 5-6 sets of samples so as to provide transport data for 30-40 nuclei.
The passive nuclear import of FITC-dextran molecules having an absorption band around 488 nm is captured using channel 1 of the confocal microscope. Figure 8 shows a representative time-lapse confocal image of the scanning area acquired in the passive channel of the microscope. The images show the central plane of the nucleus captured at six different time intervals starting with time zero, the time at which the cargo is added. In the image, the white color represents the fluorescence from the FITC-dextran. The cytoplasm remains white in color throughout the experiment with nearly constant intensity and it is assumed that the concentration of dextran in the import mixture outside the nucleus remains the same. It can be clearly seen that the fluorescence intensity inside the nucleus increases steadily with time corresponding to the entry of dextran molecules to the nucleus. The fluorescence intensity saturates at about 100 seconds after the addition of the dextran indicating an equilibrium being reached. The dark spots that can be noticed inside the nucleus are the nucleolus. It may be noted the nucleolus remains dark throughout the experiment indicating that the dextran molecules do not penetrate and enter the nucleolus.
Fig. 8. Time lapsed confocal images of passive nuclear transport of FITC labelled 10 KDa dextran molecules across HeLa cell nucleus captured using channel 1. Images of nuclei in the initial frames appear dark because a significant amount of dye-labeled dextran has not entered the nuclei. As time progresses the nuclei are appearing bright indicating the entry of fluorescently labeled dextran molecules.
To quantify the nuclear import of the molecules we determined the fluorescence intensity inside the nucleus from the images acquired at 2.2 seconds apart. This intensity is proportional to the amount of cargo that has entered the nucleus. Figure 9(a) shows a plot of the integrated nuclear fluorescence intensity F(t) with respect to time for the passive diffusion of FITC-dextran molecules in one of the nuclei. The diffusion of FITC-dextran molecules depends on the concentration gradient and at any instant of time, the rate of change of the concentration of molecules inside the nucleus is proportional to the concentration difference between nucleus and cytoplasm at the instant.
where ${C_i}(t )$ represents the concentration of molecule inside the nucleus and ${C_o}(t$) represents the concentration of molecules outside of the nucleus, which can be treated as a constant. Upon integration, this results in the first-order kinetic equation of transport for the nuclear fluorescence. The measured fluorescence intensity in channel 1 is proportional to the concentration of FITC-dextran. Hence the integrated fluorescence intensity inside the nucleus F(t) varies according to the relation [25,39] where Fmax is the nuclear fluorescence at the endpoint of the scan and k is the first-order rate constant. The solid line in Fig. 9(a) is a fit of the data to the above equation. It should be noted here that a slight variation in the nature of the graph and a spread in the rate constant and saturation intensity are observed for data obtained from different nuclei.Active nuclear import of GNP-CY3-NLS is monitored in channel 2 of the microscope using 543 nm excitation wavelength. The time-lapse images of the nuclear fluorescence captured in this channel of the microscope are shown in Fig. 10. As described above, channel 2 of the confocal microscope records only the fluorescence from CY3-labelled gold nanoparticles. Thus the images in Fig. 10 show the active nuclear transport of gold nanoparticles across the HeLa cell nuclear membrane taking place in parallel to the passive transport of FITC-dextran. The figure shows that the concentration of gold nanoparticles inside the nucleus increases steadily until it reaches a near saturation at ∼ 100 s time. The bright spots inside the nucleus indicate that the gold nanoparticles penetrate nucleolus and other sub-nuclear organelles. It can also be noted here, as is common in the active transport process, that the fluorescence intensity inside the nucleus at the end of the scan is higher than that is outside. This indicates that the rate of diffusion of molecules into the nucleus is higher than the outward diffusion rate. Figure 9(b) shows a representative graph of the nuclear fluorescence intensity of the actively transported gold nanoparticles plotted against time along with a fit to the first-order kinetic equation. Figures 8–10, the simultaneously acquired time-lapse fluorescence images of passively and actively transported model molecules and the time-dependent increase in fluorescence intensity inside the nucleus corresponding to the entry of cargo clearly show that both active and passive nuclear translocations take place at the same time. It should be noted that here we have added both the active and passive transporting molecules to the cytoplasm together for observing the simultaneous transport. The studies show that under these conditions the transport rates of passive nuclear transport are in the same range as observed in the case of passive transport alone [24]. Further studies on different systems and a large number of nuclei are required to quantify the rate constants of transport, but these preliminary studies indicate that both these transport pathways do exist together.
Fig. 9. (a) The integrated nuclear fluorescence of passive diffusion of FITC labeled dextran molecule is plotted against time. (b) The integrated fluorescence of active diffusion of CY3 labelled NLS tagged 20 nm GNP is plotted against time. The curve is fitted with a first-order kinetic equation $F(t) = {F_{\max }}(1 - \textrm{exp} ( - kt))$, where F(t) is the nuclear fluorescence at time t, Fmax is the endpoint fluorescence of the nuclear transport and k is the first-order rate constant.
Fig. 10. Time lapsed confocal images of active nuclear transport of CY3 labelled NLS tagged 20 nm GNP across HeLa cell nucleus captured using channel 2. The nuclei in the images are appearing significantly brighter compared to the background indicating the active nuclear entry of the gold nanoparticles.
4. Summary and conclusion
In summary, we have successfully designed and developed a two-channel confocal fluorescence microscope using a standard inverted microscope and two low-power lasers available in the laboratory. We successfully demonstrate the imaging capability of the microscope by imaging z-cross sections of pollen grains and standard biological cells. The microscope has a flexible design and is constructed using components normally available and can be easily reconfigured for specific applications. The image quality and resolution of the microscope are comparable to commercial microscopes and the total cost of building the microscope is only a fraction of the cost of a commercial microscope. The microscope is used to visualize the active and passive import of dye-labeled dextran molecules and gold nanoparticles into the nucleus simultaneously. The preliminary studies indicate that both the active and passive transport pathways coexist together. Experiments on a larger number of cells are currently in progress to quantify the rate constants of transport and the present report shows that the in-house constructed dual channel confocal fluorescence microscope is well suited for biomolecular transport studies.
Funding
Science and Engineering Research Board (SERB EMR/2016/003687).
Acknowledgments
Authors are grateful to Dr. Toby Joseph, Department of Physics, Birla Institute of Technology and Science, Pilani K. K. Birla Goa Campus, Goa, India – 403726, for useful discussions. Authors acknowledge the financial support from the Science and Engineering Research Board (SERB EMR/2016/003687), Government of India. P.K. Shakhi acknowledges the Council of Scientific and Industrial Research, Govt. of India for a Senior Research Fellowship. Dr. Geetha K. Varier would like to thank the Department of Science and Technology for financial support under DST-WOS-A scheme (WOS-A/PM-32/2018). We also acknowledge the support to the Department of Physics, BITS Pilani Goa campus by the Department of Science and Technology, Govt. of India under the DST-FIST scheme (Ref No. SR/FST/PSI-142/2009; SR/FST/PS-I/2017/21).
Disclosures
The authors declare no conflicts of interest.
Data availability
Data underlying the results presented in this paper are not publicly available at this time but may be obtained from the authors upon reasonable request.
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