Introduction to Widefield Microscopy

June 29, 2017

One of the most basic microscopy techniques is known as ‘Widefield Microscopy’. It is fundamentally any technique in which the entire specimen of interest is exposed to the light source with the resulting image being viewed either by the observer or a camera (which can also be attached to a computer monitor).

This article will set out to explain the differences between widefield (WF) and confocal microscopy looking especially at the differences between the imaging and illumination between the two systems. Microscope configurations for WF imaging will also be discussed looking at light paths involved as well as the problems of out-of-focus light. The more advanced WF techniques such as WF Super-Resolution will be considered below.

Basic comparison between widefield and confocal microscopy

In WF microscopy, the whole specimen on the microscope stage will be exposed to a light source (s. Figure 1). The most basic form of widefield microscopy is ‘brightfield microscopy’ in which the entire specimen is illuminated by white light either from above (in an inverted configuration), or below (in a standard upright microscope).

In a confocal laser scanning microscope, the light source for excitation of fluorescent dyes and proteins comes from laser units which are an integral part of the whole confocal system. The main advantages of confocal microscopy are that user-defined regions of interest can be selected negating the need for the entire specimen to be exposed to a fluorescent light source. In addition, the confocal microscope can be used to obtain optical sections through a specimen which can have the advantage of excluding much of the out-of-focus or background fluorescence.

Standard WF microscopes are less complex than confocal microscopes usually comprising of a white and fluorescence light source, microscope and camera (with or without an attached computer). In a confocal system, the microscope itself is only one part of a configuration including laser units, a confocal ‘scan head’ (containing pinholes for excluding out of focus light and photomultiplier tubes to collect photons from the specimen) and computers for controlling multiple parameters in the system as well as image processing.

Figure 1: Confocal vs. Widefield illumination. Confocal: Light from the light-source (Ls) is focused through a pinhole for illumination (Pi) and subsequently into the sample (S) resulting in a relatively small volume. Widefield: The whole specimen volume is exposed to light.

Comparing light sources in widefield and confocal microscopy

In a conventional laser scanning confocal microscope, around five different laser sources can be needed to cover the excitation wavelengths of commonly used fluorophores. For example, a commonly used laser is the argon-ion laser which can produce a range of excitation wavelengths which are selected by filters. The argon-ion lasers cover the green wavelengths of the excitation spectrum and are used to excite fluorophores such as FITC (fluorescein isothiocyanate). The yellow to red wavelengths of the excitation spectrum are covered by a helium-neon laser in which the range extends from approximately 543 to 632 nm. This spectrum is used to excite fluorophores such as Texas Red and rhodamine.

Before light emitting diodes (LED) were introduced as fluorescence light sources for WF microscopy, the main sources of excitation light were gas arc-lamps and these are still widely used today. The two arc-lamps which are commonly found in WF microscopes are the mercury arc-lamp (also referred to as a ‘mercury burner’ or ‘mercury vapour lamp’) and the xenon arc-lamp. The mercury arc-lamp provides excitation wavelengths across much of the visible spectrum (s. Figure 2), however, this illumination is not uniform and the main peaks are within the near-ultraviolet (UV) wavelengths (313, 334, 365, 405, 436 nm) with two other peaks in the green/yellow part of the spectrum at 546 and 579 nm.

Figure 2: Microscopy illumination is achieved with the help of certain light sources. Lasers, gas-arc lamps and LEDs all have their pros and cons and can differ distinctly in terms of their emission throughout the visible light spectrum.

Compared to mercury arc-lamps, xenon arc-lamps provide excitation wavelengths across most of the visible spectrum, but the peaks within this range do not reach the intensity of mercury burners. Although xenon arc-lamps do not extend as far into the UV part of the spectrum compared to mercury burners, their excitation range is shifted further into the infra-red wavelengths.

Although these lamps provide extremely intense light sources for fluorescence microscopy, they are not without inherent problems. The life time of these bulbs is limited with a mercury burner lasting typically 200 to 300 hours and a xenon arc-lamp lasting between 400 and 600 hours. Because they have restricted lifetimes, a careful note of the hours used should be kept with the microscope (although some systems have a built-in recorder of hours used). If gas arc-lamps are used out with their recommended lifetime range, there is a danger of explosion of the tubes. Furthermore, if the lamps are regularly switched on/off, this can significantly reduce the lifetime of the bulbs so this needs to be taken into consideration. Used arc-lamps need to be disposed of carefully and should be done so according to laboratory respectively institute regulations. Replacing and alignment of such lamps is covered in these two tutorials in the Leica Science Lab (s. Figure 3):

Figure 3: A mercury bulb is disassembled from the lamp house (background) for exchange.

Despite some of the drawbacks highlighted above, the mercury arc-lamp is still considered to be a fundamental light source for WF fluorescence microscopy due to the intensity of light produced.

The new generation of LED light sources for microscopy provide not only a full spectrum of excitation wavelengths (from around 365 to 770 nm), but also provide an intensity comparable to arc-lamps. A major advantage over the arc-lamps is the lifetime of LED sources which can be up to 50,000 hours with no warm up or cool down periods required. This also saves time as an LED unit needs only one alignment adjustment when initially fitted. Finally, waste heat is a problem with arc-lamps and they are consequently fitted in special housing units next to a microscope. As most of the electrical input with an LED is converted to light, they produce practically no waste heat.

Comparing image capture in widefield and confocal microscopy

As highlighted above, the confocal scan head contains an array of photomultiplier tubes (PMT’s) for the collection of photons from the sample. Typically, a confocal scan head will contain at least three PMT’s which are responsible for collecting red, green and blue light, but additional PMT’s are common for the collection of transmitted or reflected light. The PMT’s are not cameras as such, but are comprised of vacuum tubes which have a photon entry window at one end and an electron multiplying component in the body of the tube (s. Figure 4). The amount of photons collected is converted to an electrical signal before the image is finally assembled and displayed. Many confocal systems are also equipped with cameras similar to those used for WF microscopy.

Image capture in WF microscopy is facilitated by a more conventional camera based on photodiodes (s. Figure 4). Digital microscopy cameras contain semi-conductor detectors and the most common sensors are the Charge Coupled Devices (CCD), the Complementary Metal Oxide Semiconductors (CMOS), and the sCMOS (scientific CMOS). There are many similarities between CCD and CMOS cameras, although they process the image signal in different ways which can consequently affect the capture time of the camera as well as factors such as signal-to-noise ratio. The frame-rate of a CCD based camera can be 10-times slower than a CMOS sensor due to the intrinsic nature in their function. Choice of camera is therefore dependant on the dynamic nature of the specimen of interest. 

Figure 4: In confocal microscopy, photons are collected by photomultipliers (left). They are comprised of vacuum tubes which have a photon entry window at one end and an electron multiplying component for amplification. Microscopy camera chips consist of millions of photodiodes (right). The main composite of a photo-sensitive diode is photo-electrical silicon. Incoming photon energy is used to excite electrons of the silicon which are in turn collected in a storage well and afterwards transferred to an amplifier.

Configurations in widefield microscopy

Widefield microscopes are either upright, in which the slide is illuminated from below, or inverted, in which the slide is illuminated from above, as in Figure 5. This configuration has influence on the specimen which can be investigated. Upright microscopes are commonly used for applications with fixed samples, mounted on glass slides. Inverted microscopes have been invented to watch living cells. They are commonly grown in liquid solutions. Only a configuration with the objective below and the condenser above the specimen guarantees a close enough proximity of the objective to the specimen.

Figure 5: An upright microscope features the objective above and the condenser below the specimen (left). On an inverted microscope this set-up is turned around (right).

Epi-fluorescence is the term used to describe fluorescence microscopy in which the same lens (objective) is used to both illuminate and collect the emitted light from a sample. Light travels from the light source, passes through a filter set, through a dichromatic mirror and the shorter wavelength excitation light is reflected onto the sample via an objective. The longer wavelength emitted light from the sample travels back through the objective and passes through the dichromatic mirror to be imaged down the eyepieces and/or collected by a camera.

Trans-illumination techniques include the contrast methods such as phase contrast and differential interference contrast (DIC) microscopy. However, trans-fluorescence microscopy is a method which is neither well-known nor well-used. This method negates the use of dichromatic lenses and instead, the excitation light passes through the condenser and specimen and the emission light is collected by the objective. However, the technique had many initial problems such as high levels of background and difficulties in optically matching the condenser and objectives [1]. Recent advances which overcome these problems have led trans-fluorescence imaging to be used in areas such as in vivo imaging and in dental research [2]

Contrast methods in widefield microscopy

Contrast methods of microscopy including differential interference contrast (DIC) and phase contrast microscopy can be used in conjunction with WF fluorescence microscopy (s. Figure 6).

Combining WF fluorescence microscopy and contrast techniques can provide valuable information and images of viability and morphology. A confocal system has the ability to simultaneously capture both contrast and fluorescence images which can be overlaid using image analysis software. However, in WF fluorescence microscopy, simultaneous imaging such as this requires the use of a split view camera adaptor or the use of two separate cameras, one for contrast and one for fluorescence. More usually, a fluorescence image will be captured by the camera followed by a change in optics to contrast illumination and then the same camera will capture the contrast image. These images can then be combined using image analysis software. 

Figure 6: Neuronal cell culture in DIC (left), Phase contrast (middle) and fluorescence microscopy (right).

Resolution in widefield microscopy

The resolution of a microscope is simply the ability to distinguish detail in a specimen. It is the minimum distance which two distinct points of a specimen can still be viewed as separate entities. For more on the concepts and factors which determine resolution, please read here.

As with all conventional optical microscope systems (including confocal), resolution is determined by the numerical aperture (NA) and the wavelength of light. In light microscopy, the limit of resolution is around 200 nm. In a perfectly aligned microscope system with the highest NA optics, the limit of resolution is roughly half of the wavelength of light used to image the specimen (or excite the fluorophores). In comparing WF to confocal microscopy, the theoretical limits of resolution are similar in both methods.

However, the actual resolution achieved by a WF microscope is exacerbated by background fluorescence. As highlighted above, the entire specimen is illuminated under a WF microscope and therefore regions above and below the focal plane will also fluoresce and be captured by a camera and seen by the viewer. As such, emission wavelengths from a fluorophore can be somewhat masked by this background fluorescence leading to an overall decrease in the signal-to-noise ratio and a subsequent decrease in achievable resolution and contrast.

In a confocal system, the pinhole (within the scan head) is used to block any out-of-focus light from reaching the PMT detectors. Although this has the advantage of producing a sharply focussed image by blocking background and auto-fluorescence, some of the out-of-focus photons may have originated from the focal plane. This means that some of the information could potentially be lost due to the pinhole. There is a balance between the sharp images produced on a confocal system and inadvertently collecting out-of-focus light with a WF microscope. However, using a post-acquisition process known as ‘deconvolution’ this can resolve the blurring and focus (s. Figure 7). This computational process can re-assign photons to their point of origin.

Figure 7: Widefield fluorescence microscopy of Paramecium spec. before (left) and after (right) deconvolution.

Extending the field: a brief look at WF FRAP and Super-Resolution techniques

Fluorescence recovery after photobleaching (FRAP) is a technique which is used to examine the mobility of molecules over a period of time. This technique was initially used to study the dynamics and fluidity of lipid molecules in cellular membranes, but it can also be used to study proteins within organelles or the cytoplasm. In a FRAP experiment, a fluorescent probe is covalently attached to a molecule of interest or the target protein is engineered to express a fluorophore such as green fluorescent protein (GFP).

Once the fluorophore of interest has been localised, a target region within the cell is deliberately (and irreversibly) photobleached to erase all of the fluorescence. Over time, the bleached area will gradually become fluorescent again as more tagged target proteins move into the region whether passively (by diffusion) or actively (by transport). Such experiments can yield useful results in determining real-time movements and properties of proteins with the cellular environment (s. Figure 8). 

Figure 8: FRAP experiments can give an insight into molecule dynamics. A defined area of fluorescent molecules can be irreversibly bleached with a laser. Subsequent repopulation reveals details about the molecules’ diffusion respectively transport properties.

Although FRAP experiments were previously carried out using a confocal microscope, there are also WF solutions nowadays. Compared to the conventional confocal-based FRAP experiments, WF FRAP devices can offer higher imaging speeds with state-of-the-art cameras and also protect cells and regions of interest from the excessive photo-stress caused by confocal systems.

Super-resolution microscopy techniques are those which can image beyond the resolution limit of around 200 nm. Many super-resolution techniques have been developed over recent years and these include:

  • Single molecule localisation techniques in which small proportions of a fluorophore signal are collected over time to build up a final image. As the name suggests, it is possible to capture and image single molecules if the experimental conditions allow.
  • Structured Illumination Microscopy (SIM). In this technique, patterns are created in the phase and amplitude of WF light. The sample being imaged will also create similar patterns and the interference between the generated and sample-based patterns is used to construct intra-cellular structure.
  • Stimulated Emission Depletion (STED) Microscopy. This confocal technique requires two lasers- one for excitation of the fluorophore and another circular shaped laser beam to ‘deplete’ the emission from the same fluorophore resulting in a smaller region of image capture.

One of the single molecule localisation techniques which is achievable with WF microscopy is known as Direct Stochastic Optical Reconstruction Microscopy (dSTORM) or also known as Ground State Depletion followed by Individual Molecule Return Microscopy (GSDIM).

GSDIM/dSTORM utilises lasers and photo-switchable fluorescent probes which are switched off and on to build up an image over time. The majority of the fluorescent probes are induced into a ‘dark-state’ energy level at which they cannot emit photons. This leaves only single and well-separated fluorophores which can be located with nanometre precision.