This study was supported by JSPS KAKENHI Grant Numbers JP19H03202 to A.M., JP18H05528 and JP17H03636 to T.H., and JP17H01444 and JP18H05533 to H.Y. L.S. acknowledges the support by the Welcome Trust Strategic Awards 091911 and 107457/Z/15/Z funding advanced imaging at Micron Oxford.

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The authors have nothing to disclose.

The procedure for chromatic correction is a trade-off between accuracy and effort. To save needless efforts, it is better to know how much accuracy is required for your study. The highest accuracy may not be required for conventional wide-field (live) imaging, and thus, bright field reference images are often sufficient to correct the chromatic shift. Similarly, when the imaging condition and environment is constant, repeated use of a biological calibration will save time. On the other hand, if a highly accurate registration is desired, high-quality crosstalk or biological calibration reference images are necessary. For the best performance, reference images should be obtained with as similar conditions and timings as the target images as possible. As long as both reference and target images are obtained by the same microscopy, higher spatial resolution will improve the correction accuracy. If deconvolution is available for both reference and target images, then implementing this before correction may improve the correction accuracy. Also, for the best performance, the sampling theorem for the optical (Z) axis should be fulfilled in both the reference and target file for precise subpixel interpolation (protocol step 2.1.3).
Failure to correct chromatic shift leads to incorrect conclusions. Furthermore, using the wrong calibration may even worsen the chromatic shifts rather than correcting them, and this therefore needs to be avoided. We have summarized the possible causes of failures, and their common solutions, in Table 2. To examine the cause of a failure, in the first place, it is necessary to visually check if the chromatic shift in the reference image is precisely corrected (protocol step 3.12). Most failures are due to the quality of the reference images and are easily remedied as per the descriptions in Table 2. Regarding to the quality of reference images, it is important to note that the accuracy of global alignment decreases if the entire field of view is not filled with the sample (Figure 7, Table 2). Compared to the good example shown in Figure 7A, the bad example shown in Figure 7B contains only three nuclear envelopes in the upper-left region, and Chromagnon failed to align a part of this image. This is because the global alignment method of Chromagnon splits the field of view into four regions (Figure 7C) in order to measure the differences in rotation and magnification with high accuracy3. This method, if correctly operated, is one order more accurate than other linear methods such as the log polar transformation and simplex methods3. If any of the four regions are unavailable, then Chromagnon will switch to less effective linear methods. Therefore, for the best performance, the examples shown in Figure 7B and Figure 7C are undesirable, and the four regions should be filled with objects. Users can check if any quadratic region of the field of view is unavailable for measurement by looking at the log file (&#34;Chromagnon.log&#34;; see protocol step 3.10). Fortunately, this problem can be easily overcome by averaging multiple biological calibration images or using local alignment for crosstalk or bright-field reference images (Table 2). Contrary to the case of failure to correct reference images, failure to correct target images is more difficult to identify. Because such failures arise due to differences in file formats, imaging conditions, imaging timings, imaging/alignment methods between the reference and target images (Table 2), users should always be careful when using reference images that are obtained in different conditions/timings from the target images. Some example images are available for testing (<a href="https://github.com/macronucleus/Chromagnon" target="_blank">https://github.com/macronucleus/Chromagnon</a>) to obtain concrete idea of the good and bad example images.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Problem</td>
			<td>Cause</td>
			<td>Solution</td>
		</tr>
		<tr>
			<td>Failed to correct the reference image</td>
			<td>Low contrast</td>
			<td>Acquire a higher contrast image if possible. If a bright-field reference image is used, reacquire the image in a water-based solution to obtain higher contrast of the cell. Alternatively, try applying computational noise reduction (e.g. Gaussian filtering). Turn off local alignment, which is more sensitive to noise.</td>
		</tr>
		<tr>
			<td></td>
			<td>Contamination of unrelated images</td>
			<td>Remove the source of the unrelated images in the sample if possible. For crosstalk reference images, check the excitation spectra of the dyes used for the target images. If the dyes are excited during acquisition of a crosstalk image (e.g. Alexa Fluor 568 or 594), consider other dyes (e.g. Alexa Fluor 555). If dusts on the camera chip creates an obvious channel difference, clean the camera chip or use a computational flat-fielding method.</td>
		</tr>
		<tr>
			<td></td>
			<td>An extremely bright spot made by a cosmic ray</td>
			<td>Acquire the image again if possible. Alternatively, try applying computational noise reduction (e.g. Median or Gaussian filtering).</td>
		</tr>
		<tr>
			<td></td>
			<td>Deconvolution artifacts (artificial signals at the axial and lateral edges)</td>
			<td>Trim the edge pixels or Z sections after deconvolution. If one side is trimmed, the other side should also be trimmed to maintain the image center.</td>
		</tr>
		<tr>
			<td></td>
			<td>The Z step size too sparse</td>
			<td>A Z stack should be acquired to fulfill the Nyquist criterion as written in protocol 2.1.3.</td>
		</tr>
		<tr>
			<td></td>
			<td>Optical aberration</td>
			<td>Spherical aberration is the major aberration caused by users. Choose the right objective lens for the sample and use a coverslip thickness of 170 &#181;m. If the objective lens is equipped with a correction ring, adjust it to find the position where the highest fluorescence count is obtained from the focus. In the case of an oil-immersion objective without a correction ring, adjust the refractive index of the immersion oil that increases the fluorescence count at the focus.</td>
		</tr>
		<tr>
			<td></td>
			<td>Field of view is unfilled (Fig. 7)</td>
			<td>In the case of biological calibration reference images, average many images. In the case of crosstalk or bright-field reference images, use local alignment.</td>
		</tr>
		<tr>
			<td></td>
			<td>An unidentified software bug</td>
			<td>Report the issue through GitHub (<a href="https://github.com/macronucleus/Chromagnon/issues" target="_blank">https://github.com/macronucleus/Chromagnon/issues)</a></td>
		</tr>
		<tr>
			<td>Failed to correct the target image</td>
			<td>Metadata of the image file is lost</td>
			<td>Use the original microscope file format which contains complete metadata, and avoid converting to a multipage tiff file before processing. Use the same ordering of channels as written in protocol 3.3.</td>
		</tr>
		<tr>
			<td></td>
			<td>Wrong alignment methods for the given microscopy</td>
			<td>Do not apply the local alignment method when measuring from biological calibration reference images to target images. Do not use crosstalk reference images other than wide-field microscopy.</td>
		</tr>
		<tr>
			<td></td>
			<td>Differences in imaging conditions</td>
			<td>Keep the imaging conditions constant between the reference and the target images as written in protocol 2.3.3.</td>
		</tr>
		<tr>
			<td></td>
			<td>Differences in sample (including coverslip)</td>
			<td>Always use the same mounting medium, coverslip (e.g. No. 1.5H) and a similar depth of focus.</td>
		</tr>
		<tr>
			<td></td>
			<td>Microscope drift since the calibration was last made</td>
			<td>Make a calibration as often as every two weeks. Keep the temperature constant, and use a floating table to avoid hardware drift of the microscope.</td>
		</tr>
	</tbody>
</table>
Table 2: Troubleshooting for chromatic correction.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/60800/60800fig07.jpg" /> 
Figure 7: Examples of reference images. Nuclear envelope in fission yeast cells labeled with GFP and mCherry. Images were acquired with conventional wide-field microscopy. Chromatic shifts were corrected using Chromagnon without local alignment using the images themselves as reference images. Images were then deconvolved to show the details. (A) A good example with many objects in the field of view. (B) A bad example with objects only at the top-left corner. Misalignment is obvious at a certain region of the image. (C) An undesirable example where one of the quadrisection (separated by dotted cross lines) is empty. Scale bar in panel A indicates 5 &#181;m for the full field view and 1.25 &#181;m for the enlarged view and is applicable to all panels. <a href="https://www.jove.com/files/ftp_upload/60800/60800fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In this protocol, we described three different reference types (Table 1). Among them, crosstalk reference images and biological calibration reference images need further careful discussion. For crosstalk reference images, samples stained with DAPI or Hoechst 33342, and mounted in glycerol or commercial mounting media can be efficiently used to align the blue, green, and red channels. Similarly, Alexa Fluor 488 can be used to align the green and red channels. However, obtaining crosstalk fluorescence is often difficult since many blue dyes except DAPI and Hoechst are dimmer and decay faster than most green and red dyes. Furthermore, the emission spectra of modern dyes are narrower, which makes the alignment of more than three channels by this method challenging. Attention should also be paid to some common red dyes (e.g., Alexa Flour 568 and 594, but not Alexa Fluor 555) that can be excited by violet light, which prevent obtaining high-contrast crosstalk images from blue dyes. Another drawback is that this method cannot measure the chromatic aberration of excitation light paths in multicolor excitation, because only a single excitation wavelength is used for excitation (Table 1). As most advanced microscopy uses altered illumination optics, the application of this method is limited. Still, its higher correction accuracy is sufficiently advantageous for it to be described in this protocol. In general, a crosstalk image should be taken after a target image to prevent bleaching or phototoxic effects. For SMLM observed with the wide-field mode, a reference image should be acquired before acquiring a target image as fluorescence dyes can be bleached while imaging.
Biological calibration reference images allow users to easily align any desired number of channels at the cost of additional sample preparation. Another advantage of biological calibration reference images is the availability of &#34;averaging&#34; multiple references that helps fill all fields of view. This method may suffer from differences in imaging conditions if the calibration sample is prepared on a different slide. Most of this problem can be solved if both targets and references are prepared on the same slide by using commercial chambered coverglasses (Table 1), and other imaging conditions are kept constant as in protocol step 2.3.3. In this case, a correction accuracy similar to that of crosstalk reference images can be expected3. The protocol to use phalloidin as shown here is one of the easiest ways to stain a single cellular structure with multiple colors. There are numerous possible scenarios to prepare biological calibration samples. For immunostaining, a sample can be labeled with a single primary antibody followed by staining with secondary antibodies of multiple colors. In this way, a single target structure can be labeled with multiple colors. Alternatively, 5-ethynyl-2&#39;-deoxyuridine, detected by &#34;click&#34; chemistry labels newly synthesized DNA in multiple colors at high density, as described in detail previously8. For live cells, it is useful to prepare a transgenic strain harboring two copies of a gene that are fused to GFP or mCherry to label the same structure with two colors. If the copy number of the gene is critical as often observed for membrane proteins, a single copy of the gene can be tandemly fused to GFP and mCherry (Figure 7). Photoconvertible fluorescent proteins, such as mEOS218, can also be used by illuminating a moderate level of violet light to obtain both protein species with or without photoconversion. Under low oxygen conditions, GFP can also be used as a photoconvertible protein from green to red19,20. Choosing the right calibration sample will thus make the experiment more robust.

Multicolor imaging is one of the fundamental aspects of biological fluorescence microscopy, in cases where the spatial relationship of different molecules or structures is of major interest. Chromatic aberration, an optical aberration of polychromatic light caused by dispersion, changes the apparent position of the colored objects of interest. Similarly, microscopes equipped with multiple cameras devoted to acquiring each color have more complex chromatic shifts due to differences in optical elements and imperfect alignment among the channels. Thus, such chromatic shifts may lead to a false conclusion unless explicitly corrected by the user. Although chromatic shifts have not been a major problem as long as the resolution of microscopy is limited by the classical resolution limit, recent development of super-resolution microscopy1 has prompted the need for more accurate correction of chromatic shifts.
It has been a common practice to measure chromatic shifts of microscope systems using a multicolor bead calibration slide2. The bead-based calibration method is appropriate for measuring chromatic shifts from the entire optics of the microscope towards the surface of the coverslip2. This method, however, is unable to measure chromatic shifts in the biological samples of interest. It is important to note that many biological samples are three-dimensional (3D), and the chromatic shifts of such samples are different from those at the surface of the coverslip. Furthermore, chromatic shifts change with imaging conditions2,3. We have measured the chromatic shifts in 3D biological samples and found that the uncertainty of chromatic shifts was often as much as 350 nm by the classical multicolor-bead calibration method3. Therefore, chromatic shifts need to be measured in biological samples at the depth of interest and under the imaging conditions being used.
Here, we describe procedures to measure chromatic shifts in biological samples and correct these shifts using our software, Chromagnon3. To measure chromatic shifts in biological samples, our method uses two kinds of data sets, a &#34;target&#34; image and a &#34;reference&#34; image. The &#34;target&#34; image is a multicolor image of interest, for example, images stained for DNA, nuclear envelope, and microtubules. It is often impossible to measure chromatic shifts in such an image. Therefore, we need a &#34;reference&#34; image that is dedicated to measure the chromatic shifts in the sample. The only definition of a &#34;reference&#34; image is a multicolor image of the same object. In this sense, a multicolor beads image is also a type of reference image. Here, we describe three different types of reference image that are used to measure chromatic shifts in the biological samples: &#34;crosstalk reference images&#34;, &#34;bright-field reference images&#34; and &#34;biological calibration reference images&#34;. The type of reference image depends on the type of microscope being used or the correction accuracy required as summarized in Table 1.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Crosstalk</td>
			<td>Bright-field</td>
			<td>Biological calibration (on a different slide)</td>
			<td>Biological calibration (on the same slide)</td>
		</tr>
		<tr>
			<td>Accuracya</td>
			<td>+++</td>
			<td>+</td>
			<td>++b</td>
			<td>+++</td>
		</tr>
		<tr>
			<td>Simplicity</td>
			<td>++</td>
			<td>+++</td>
			<td>++</td>
			<td>++</td>
		</tr>
		<tr>
			<td>Applicable microscopy</td>
			<td>Wide-field</td>
			<td>Wide-field</td>
			<td>All</td>
			<td>All</td>
		</tr>
		<tr>
			<td>Availability of local alignment</td>
			<td>+</td>
			<td>+</td>
			<td>-c</td>
			<td>-c</td>
		</tr>
		<tr>
			<td colspan="5">a: Number of &#34;+&#34; indicates increasing rating. Single plus is about 50 nm and three plus is about 15 nm in 3D. 
			b: The accuracy depends on how much the variable imaging conditions are kept constant. 
			c: Local calibration measured by multicolor bead samples can be combined as described in protocol section 4.</td>
		</tr>
	</tbody>
</table>
Table 1: Parameters when choosing the type of reference images.
&#34;Crosstalk reference images&#34; have the highest correction accuracy and are relatively simple to accomplish3,4 (Table 1). The drawback is their limitation in microscopy applications due to their incapability of measuring chromatic shifts in excitation paths. Also, to obtain such images, the microscope should be equipped with multiband dichroic mirrors, and emission filters that are independently controlled from the excitation filters or light sources. Suitable microscopy includes conventional wide-field microscopy, single molecule localization microscopy (SMLM) such as photo-activated localization microscopy/stochastic optical reconstruction microscopy (PALM/STORM)5,6 and expansion microscopy7 observed with wide-field microscopy. A crosstalk reference image is acquired from the target sample itself. It is an image of crosstalk (bleed-through) fluorescence of a dye obtained in all required channels. Fluorescence emission always expands towards the longer wavelengths, therefore dyes with the shortest emission wavelength are excited to obtain crosstalk fluorescence in channels of longer wavelengths. For example, when the sample is stained with blue, green, and red, only the blue dye is excited, and the emission light is obtained in the blue, green, and red channels. In this protocol, DNA stained with 4&#8242;,6-diamidino-2-phenylindole (DAPI) was used to obtain crosstalk fluorescence.
&#34;Bright-field reference images&#34; are an easier and less phototoxic alternative to &#34;crosstalk reference images&#34; but are the least accurate3 (Table 1). These are bright field images of the target sample, acquired in all the color channels used in the target image.
&#34;Biological calibration reference images&#34; have the advantage of being applicable to any type of microscopy due to their ability to measure the chromatic shifts both in the excitation and emission paths3,8 (Table 1). Suitable microscopy includes wide-field microscopy, confocal microscopy, light sheet microscopy, stimulated emission depletion (STED)9, structured illumination microscopy (SIM)10, Airyscan/SORA11,12, SMLM observed with the total internal reflection fluorescence (TIRF) mode, Olympus super resolution (OSR)13, and so forth. A biological calibration reference image is acquired from a calibration sample similarly prepared as the target sample, but with staining of a single structure with multiple colors. The correction accuracy excels the resolution of most super-resolution microscopy and preparing a biological calibration sample can be relatively simple. Another advantage is the availability to &#34;average&#34; multiple reference images. Therefore, even though the individual images contain poor information for the measurement of chromatic shifts, the information content can be increased by averaging multiple images. The accuracy depends on how much the imaging conditions are kept constant. In this regard, the best performance is obtained when both target and reference samples are on the same slide, using, for example, 8-well chambered coverglasses (Table 1, right-most). In this protocol, actin stained with three colors of phalloidin was used as a biological calibration.
Once a reference image is obtained, then the chromatic shift is measured and corrected by our software Chromagnon. There is no limitation on the number of channels, Z sections and time frames that Chromagnon can measure and correct the chromatic shifts for. Chromagnon measures chromatic shifts in two steps. The first step acquires the &#34;global&#34; or &#34;affine&#34; alignment parameters of translation in the X, Y, Z axes, magnification along the X, Y, Z axes, and rotation around the Z axis. The calculation accuracy of the global alignment is ~16 nm in 3D and ~8 nm in 2D. The second step is an optional 2D iterative &#34;local alignment&#34; on projected images to obtain a higher accuracy. In the local alignment process, the images are subdivided into multiple regions and chromatic shifts in these local regions are measured. Subsequently, the regions are further divided and chromatic shifts in the subregions are measured iteratively until the number of pixels in the region reaches the minimum number of pixels (usually 60 x 60 pixels). The resulting local alignment map is combined with the global alignment parameter and is applied to the target image by an elastic transformation. Following this step, the calculation accuracy is improved to ~14 nm in 3D and ~6 nm in 2D. The local alignment is not suitable for biological calibration reference images because biological structure in the reference is different from that in the target (Table 1). Therefore, only global alignment is used for biological calibration reference images.
The local chromatic shifts originate from two sources; microscope instrumental local distortion and biological structural inhomogeneity. Because microscope instrumental local distortion is constant, this can be measured from the multicolor beads reference sample and corrected as a fixed parameter. Chromagnon can combine the microscope instrumental local distortion map and the global alignment parameters from the biological calibrations (Table 1). Using this method, it is expected that the average accuracy of biological calibration will be improved by an additional 1&#8722;2 nm.
Here, we describe a protocol to correct the chromatic shifts of 3D fluorescence images using our software Chromagnon, from the easiest low end to the highest accuracy. We use immunostaining of HeLa cells as an example and observed them using 3D wide-field microscopy and 3D-SIM. In the first section, we describe how to prepare target samples and biological calibration samples. This part of the protocol should be optimized for the specific targets of the research. In the second section, we describe the acquisition methods for three kinds of reference images by microscopes. The assumption was to obtain blue, green, and red channels but channel composition should be modified by the specific targets of the research and by the setups of the microscope. It does not matter if the microscope is equipped with a single camera or multiple cameras. In the third section, we describe how one can use our software to measure and correct chromatic shifts of the target image by using reference images. Finally, in the fourth section, we describe a method to complement the biological calibration reference images by using a microscope&#39;s instrumental local calibration.

<table><tbody><tr><td>16% formaldehyde solution</td><td>Polyscience</td><td>18814-10</td><td></td></tr><tr><td>35 mm glass-bottom dish</td><td>MatTek</td><td>P35G-1.5-10-C</td><td></td></tr><tr><td>Alexa Fluor 405 phalloidin</td><td>Thermo Fisher Scientific</td><td>A30104</td><td></td></tr><tr><td>Alexa Fluor 488 phalloidin</td><td>Thermo Fisher Scientific</td><td>A12379</td><td></td></tr><tr><td>Alexa Fluor 594 phalloidin</td><td>Thermo Fisher Scientific</td><td>A12381</td><td></td></tr><tr><td>Bovine Serum</td><td>Thermo Fisher Scientific</td><td>16170078</td><td></td></tr><tr><td>Coverslip</td><td>Matsunami</td><td>No. 1S HT</td><td></td></tr><tr><td>DAPI (4&#039;,6-Diamidino-2-Phenylindole, Dihydrochloride)</td><td>Thermo Fisher Scientific</td><td>D1306</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Modified Eagle Medium with L-Gln and sodium pyruvate</td><td>Nacalai Tesque</td><td>08458-16</td><td></td></tr><tr><td>Mounting medium (VECTASHIELD)</td><td>Vector Laboratories</td><td>H-1000</td><td></td></tr><tr><td>Mouse anti-tubulin monoclonal antibody (TAT1)</td><td></td><td></td><td>Described in Ref 15.</td></tr><tr><td>Nunc Lab-Tek II chambered coverglass (8 well)</td><td>Thermo Fisher Scientific</td><td>155409</td><td></td></tr><tr><td>Rabbit anti-emerin polyclonal antibody (ED1)</td><td></td><td></td><td>A gift from Hiroshi Yorifuji, Gunma University, Gunma, Japan and Kiichi Arahata, National Center of Neurology and Psychiatry, Tokyo, Japan; deceased.</td></tr><tr><td>Secondary antibody with Alexa Fluor 488</td><td>Thermo Fisher Scientific</td><td>A-11034</td><td></td></tr><tr><td>Secondary antibody with Alexa Fluor 555</td><td>Thermo Fisher Scientific</td><td>A-21424</td><td></td></tr><tr><td>Secondary antibody with Alexa Fluor 594</td><td>Thermo Fisher Scientific</td><td>A-11032</td><td></td></tr><tr><td>TetraSpeck Microspheres, 0.2 &amp;micro;m</td><td>Thermo Fisher Scientific</td><td>T7280</td><td></td></tr></tbody></table>

high-accuracy correction of 3d chromatic shifts in the age of super-resolution biological imaging using chromagnon

Quantitative multicolor fluorescence microscopy relies on the careful spatial matching of color channels acquired at different wavelengths. Due to chromatic aberration and the imperfect alignment of cameras, images acquired in each channel may be shifted, and magnified, as well as rotated relative to each other in any of the three dimensions. With the classical calibration method, chromatic shifts are measured by multicolor beads attached to the surface of a coverslip, and a number of software are available to measure the chromatic shifts from such calibration samples. However, chromatic aberration can vary with depth, change with observation conditions and be induced by the biological sample itself, thus hindering determination of the true amount of chromatic shift in the sample of interest and across the volume. Correcting chromatic shifts at higher accuracy is particularly relevant for super-resolution microscopy where only slight chromatic shifts may affect quantitative analyses and alter the interpretation of multicolor images. We have developed an open-source software Chromagnon and accompanying methods to measure and correct 3D chromatic shifts in biological samples. Here we provide a detailed application protocol that includes special requirements for sample preparation, data acquisition, and software processing to measure chromatic shifts in biological samples of interest.

An example of chromatic shift correction using a crosstalk reference image is shown in Figure 5. The image was obtained with a wide-field microscope equipped with a single camera. Fluorescence emission from DAPI was used as a reference (Figure 5A,B) to correct the blue, green, and red channels. The image comprises 3 channels of 60 Z slices, each composed of 256 x 256 pixels. The images were deconvolved before measuring the chromatic shifts. Measuring the local chromatic shifts using Chromagnon took 51 s on a Mac with Intel Core i7 (quad core, 8-threads, 2.7 GHz), 16 GB RAM and 1 TB flash storage. The alignment parameter was applied to the target image (Figure 5C,D), which has exactly the same number of voxels as the reference image. Preparing the aligned file took 3 s. As a result of trimming the edge pixels during the alignment process (crop margins checkbox in Figure 1C), the number of voxels was reduced after alignment (Figure 5B, 51 Z slices, 252 x 251 pixels). DNA in the anaphase bridge (indicated by arrowheads) is seen incorrectly outside of the nuclear envelope before alignment (Figure 5C, obvious in the bottom panel showing the XZ view), but as expected inside the envelope after alignment (Figure 5D).
An example of chromatic shift correction using a biological calibration reference image is shown in Figure 6. The images were obtained with a SIM microscope equipped with three cameras. Three images of HeLa cells stained with phalloidins conjugated to blue, green, and red dyes were averaged (Figure 6A). The reference image comprises 3 channels of 76 Z slices, each composed of 1,024 x 1,024 pixels. Measuring chromatic shifts using Chromagnon without local alignment required 194 s on the Mac system described above. The parameter was applied to a target image consisting of 3 channels of 73 Z slices, each composed of 1,024 x 1,024 pixels. Generation of the aligned file took 25 s. The XZ view shows incorrect channel positions along the Z, and slightly along the X directions (Figure 6A,C) but this misregistration was corrected after alignment (Figure 6B,D).
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60800/60800fig05.jpg" /> 
Figure 5: An example of alignment with a crosstalk reference image. HeLa cells were stained with DAPI for DNA (shown in magenta), Alexa Fluor 488 (shown in yellow) for nuclear envelope, and Alexa Fluor 555 (shown in blue) for microtubules. Images were acquired by 3D wide-field microscopy with a single camera and deconvolved. (A,B) A representative crosstalk reference image using DAPI emission, before and after alignment by Chromagnon. Three color channels are shown as overlaid. (C,D) An optical section of the 3D stack in three channels before and after alignment. Axial chromatic aberration is obvious at the anaphase bridge shown by arrowheads. Scale bar in panel A indicates 5 &#181;m for all panels. <a href="https://www.jove.com/files/ftp_upload/60800/60800fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60800/60800fig06.jpg" /> 
Figure 6: An example of alignment with a biological calibration reference image Images were acquired with 3D-SIM equipped with three cameras. (A,B) A reference image averaged from three images before (A) and after (B) alignment. HeLa cells were stained with phalloidin conjugated with Alexa Fluor 405, 488 or 594. (C,D) The target image before (C) and after (D) alignment. HeLa cells were stained with DAPI for DNA (shown in magenta), Alexa Fluor 488 (shown in yellow) for nuclear envelope, and Alexa Fluor 594 (shown in blue) for microtubules. Scale bar in panel A indicates 5 &#181;m for all panels. <a href="https://www.jove.com/files/ftp_upload/60800/60800fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about High-Accuracy Correction of 3D Chromatic Shifts in the Age of Super-Resolution Biological Imaging Using Chromagnon at JoVE.com

High-Accuracy Correction of 3D Chromatic Shifts in the Age of Super-Resolution Biological Imaging Using Chromagnon

Correction of chromatic shifts in three-dimensional (3D) multicolor fluorescence microscopy images is crucial for quantitative data analyses. This protocol is developed to measure and correct chromatic shifts in biological samples through acquisition of suitable reference images and processing with the open-source software, Chromagnon.

High-Accuracy Correction of 3D Chromatic Shifts in the Age of Super-Resolution Biological Imaging Using Chromagnon

Quantitative multicolor fluorescence microscopy relies on the careful spatial matching of color channels acquired at different wavelengths. Due to ...

high-accuracy-correction-3d-chromatic-shifts-age-super-resolution

Research

JoVE Journal

Biology

7.4K Views.  National Institute of Information and Communications Technology. Marsh koraal microscopie beelden bevatten Chromatic Shift in drie dimensies. Dat schaadt de colocalisatie-analyse van beelden met hoge resolutie. Het is over het algemeen moeilijk om Chromatic Shift afkomstig van een biologisch monster zelf te verwijderen, onze methode kan chromatische verschuiving in een biologisch monster corrigeren met behulp van afzonderlijk verworven referentiebeelden.Het aantonen van de procedure zal Takako Koujin een technicus van mijn groep. Begin met het zaaie...

Video: High-Accuracy Correction of 3D Chromatic Shifts in the Age of Super-Resolution Biological Imaging Using Chromagnon

Sample Drift Correction Following 4D Confocal Time-lapse Imaging

The generation of four-dimensional (4D) confocal datasets; consisting of 3D image sequences over time; provides an excellent methodology to capture cellular behaviors involved in developmental processes.&#160; The ability to track and follow cell movements is limited by sample movements that occur due to drift of the sample or, in some cases, growth during image acquisition. Tracking cells in datasets affected by drift and/or growth will incorporate these movements into any analysis of cell position. This may result in the apparent movement of static structures within the sample. Therefore prior to cell tracking, any sample drift should be corrected. Using the open source Fiji distribution 1 &#160;of ImageJ 2,3 and the incorporated LOCI tools 4, we developed the Correct 3D drift plug-in to remove erroneous sample movement in confocal datasets. This protocol effectively compensates for sample translation or alterations in focal position by utilizing phase correlation to register each time-point of a four-dimensional confocal datasets while maintaining the ability to visualize and measure cell movements over extended time-lapse experiments.

Time-lapse microscopy allows the visualization of developmental processes. Growth or drift of samples during image acquisition reduces the ability to accurately follow and measure cell movements during development. We describe the use of open source image processing software to correct for three dimensional sample drift over time.

The generation of four-dimensional (4D) confocal datasets; consisting of 3D image sequences over time; provides an excellent methodology to capture ...

Bioengineering

Highly Resolved Intravital Striped-illumination Microscopy of Germinal Centers

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick tissue samples and organs in health and disease. By controlling the scanning pattern in multi-photon microscopy and applying appropriate numerical algorithms, we developed a striped-illumination approach, which enabled us to achieve 3-fold better axial resolution and improved signal-to-noise ratio, i.e. contrast, in more than 100 &#181;m tissue depth within highly scattering tissue of lymphoid organs as compared to standard multi-photon microscopy. The acquisition speed as well as photobleaching and photodamage effects were similar to standard photo-multiplier-based technique, whereas the imaging depth was slightly lower due to the use of field detectors. By using the striped-illumination approach, we are able to observe the dynamics of immune complex deposits on secondary follicular dendritic cells &#8211; on the level of a few protein molecules in germinal centers.

High-resolution intravital imaging with enhanced contrast up to 120 &#181;m depth in lymph nodes of adult mice is achieved by spatially modulating the excitation pattern of a multi-focal two-photon microscope. In 100 &#181;m depth we measured resolutions of 487 nm (lateral) and 551 nm (axial), thus circumventing scattering and diffraction limits.

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick ...

Immunology and Infection

Excitation-Scanning Hyperspectral Imaging Microscopy to Efficiently Discriminate Fluorescence Signals

Several techniques rely on detection of fluorescence signals to identify or study phenomena or to elucidate functions. Separation of these fluorescence signals were proven cumbersome until the advent of hyperspectral imaging, in which fluorescence sources can be separated from each other as well as from background signals and autofluorescence (given knowledge of their spectral signatures). However, traditional, emission-scanning hyperspectral imaging suffers from slow acquisition times and low signal-to-noise ratios due to the necessary filtering of both excitation and emission light. It has been previously shown that excitation-scanning hyperspectral imaging reduces the necessary acquisition time while simultaneously increasing the signal-to-noise ratio of acquired data. Using commercially available equipment, this protocol describes how to assemble, calibrate, and use an excitation-scanning hyperspectral imaging microscopy system for separation of signals from several fluorescence sources in a single sample. While highly applicable to microscopic imaging of cells and tissues, this technique may also be useful for any type of experiment utilizing fluorescence in which it is possible to vary excitation wavelengths, including but not limited to: chemical imaging, environmental applications, eye care, food science, forensic science, medical science, and mineralogy.

Spectral imaging has become a reliable solution for identification and separation of multiple fluorescence signals in a single sample and can readily distinguish signals of interest from background or autofluorescence. Excitation-scanning hyperspectral imaging improves on this technique by decreasing the necessary image acquisition time while simultaneously increasing the signal-to-noise ratio.

Several techniques rely on detection of fluorescence signals to identify or study phenomena or to elucidate functions. Separation of these fluorescence ...

Engineering

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Developmental Biology

Conducting Multiple Imaging Modes with One Fluorescence Microscope

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to conventional epi-fluorescence microscopy, various imaging techniques have been developed to achieve specific experimental goals. Some of the widely used techniques include single-molecule fluorescence resonance energy transfer (smFRET), which can report conformational changes and molecular interactions with angstrom resolution, and single-molecule detection-based super-resolution (SR) imaging, which can enhance the spatial resolution approximately ten&#160;to twentyfold compared to diffraction-limited microscopy. Here we present a customer-designed integrated system, which merges multiple imaging methods in one microscope, including conventional epi-fluorescent imaging, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging. Different imaging methods can be achieved easily and reproducibly by switching optical elements. This set-up is easy to adopt by any research laboratory in biological sciences with a need for routine and diverse imaging experiments at a reduced cost and space relative to building separate microscopes for individual purposes.

Here we present a practical guide of building an integrated microscopy system, which merges conventional epi-fluorescent imaging, single-molecule detection-based super-resolution imaging, and multi-color single-molecule detection, including single-molecule fluorescence resonance energy transfer imaging, into one set-up in a cost-efficient way.

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to ...

Chromosomics: Detection of Numerical and Structural Alterations in All 24 Human Chromosomes Simultaneously Using a Novel OctoChrome FISH Assay

Fluorescence in situ hybridization (FISH) is a technique that allows specific DNA sequences to be detected on metaphase or interphase chromosomes in cell nuclei1. The technique uses DNA probes with unique sequences that hybridize to whole chromosomes or specific chromosomal regions, and serves as a powerful adjunct to classic cytogenetics. For instance, many earlier studies reported the frequent detection of increased chromosome aberrations in leukemia patients related with benzene exposure, benzene-poisoning patients, and healthy workers exposed to benzene, using classic cytogenetic analysis2. Using FISH, leukemia-specific chromosomal alterations have been observed to be elevated in apparently healthy workers exposed to benzene3-6, indicating the critical roles of cytogentic changes in benzene-induced leukemogenesis. 
Generally, a single FISH assay examines only one or a few whole chromosomes or specific loci per slide, so multiple hybridizations need to be conducted on multiple slides to cover all of the human chromosomes. Spectral karyotyping (SKY) allows visualization of the whole genome simultaneously, but the requirement for special software and equipment limits its application7. Here, we describe a novel FISH assay, OctoChrome-FISH, which can be applied for Chromosomics, which we define here as the simultaneous analysis of all 24 human chromosomes on one slide in human studies, such as chromosome-wide aneuploidy study (CWAS)8. The basis of the method, marketed by Cytocell as the Chromoprobe Multiprobe System, is an OctoChrome device that is divided into 8 squares, each of which carries three different whole chromosome painting probes (Figure 1). Each of the three probes is directly labeled with a different colored fluorophore, green (FITC), red (Texas Red), and blue (Coumarin). The arrangement of chromosome combinations on the OctoChrome device has been designed to facilitate the identification of the non-random structural chromosome alterations (translocations) found in the most common leukemias and lymphomas, for instance t(9;22), t(15;17), t(8;21), t(14;18)9. Moreover, numerical changes (aneuploidy) in chromosomes can be detected concurrently. The corresponding template slide is also divided into 8 squares onto which metaphase spreads are bound (Figure 2), and is positioned over the OctoChrome device. The probes and target DNA are denatured at high-temperature and hybridized in a humid chamber, and then all 24 human chromosomes can be visualized simultaneously. 
OctoChrome FISH is a promising technique for the clinical diagnosis of leukemia and lymphoma and for detection of aneuploidies in all chromosomes. We have applied this new Chromosomic approach in a CWAS study of benzene-exposed Chinese workers8,10.

A novel fluorescence in situ hybridization (FISH) method that simultaneously examines both numerical and structural chromosome alterations, particularly the specific chromosomal translocations associated with leukemia and lymphoma, of all 24 human chromosomes on a single device in one hybridization, is described.

Fluorescence in situ hybridization (FISH) is a technique that allows specific DNA sequences to be detected on metaphase or interphase chromosomes in cell ...

Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus

Super-resolution microscopy has been widely used to study protein interactions and subcellular structures in many organisms. In photosynthetic organisms, however, the lateral resolution of super-resolution imaging is only ~100 nm. The low resolution is mainly due to the high autofluorescence background of photosynthetic cells caused by high-intensity lasers that are required for super-resolution imaging, such as stochastic optical reconstruction microscopy (STORM). Here, we describe a photobleaching-assisted STORM method which was developed recently for imaging the marine picocyanobacterium Prochlorococcus. After photobleaching, the autofluorescence of Prochlorococcus is effectively reduced so that STORM can be performed with a lateral resolution of ~10 nm. Using this method, we acquire the in vivo three-dimensional (3-D) organization of the FtsZ protein and characterize four different FtsZ ring morphologies during the cell cycle of Prochlorococcus. The method we describe here might be adopted for the super-resolution imaging of other photosynthetic organisms.

Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus

Here we describe a photobleaching method to reduce the autofluorescence of cyanobacteria. After photobleaching, stochastic optical reconstruction microscopy is used to obtain three-dimensional super-resolution images of the cyanobacterial FtsZ ring.

Super-resolution microscopy has been widely used to study protein interactions and subcellular structures in many organisms. In photosynthetic organisms, ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, making it difficult to isolate inputs from individual sources with electrical stimulation. However, by using channelrhodopsin assisted circuit mapping (CRACM), this limitation can now be overcome. Here, we report a method to use CRACM to map ascending inputs from lower auditory brainstem nuclei and commissural inputs to an identified class of neurons in the inferior colliculus (IC), the midbrain nucleus of the auditory system. In the IC, local, commissural, ascending, and descending axons are heavily intertwined and therefore indistinguishable with electrical stimulation. By injecting a viral construct to drive expression of a channelrhodopsin in a presynaptic nucleus, followed by patch clamp recording to characterize the presence and physiology of channelrhodopsin-expressing synaptic inputs, projections from a specific source to a specific population of IC neurons can be mapped with cell type-specific accuracy. We show that this approach works with both Chronos, a blue light-activated channelrhodopsin, and ChrimsonR, a red-shifted channelrhodopsin. In contrast to previous reports from the forebrain, we find that ChrimsonR is robustly trafficked down the axons of dorsal cochlear nucleus principal neurons, indicating that ChrimsonR may be a useful tool for CRACM experiments in the brainstem. The protocol presented here includes detailed descriptions of the intracranial virus injection surgery, including stereotaxic coordinates for targeting injections to the dorsal cochlear nucleus and IC of mice, and how to combine whole cell patch clamp recording with channelrhodopsin activation to investigate long-range projections to IC neurons. Although this protocol is tailored to characterizing auditory inputs to the IC, it can be easily adapted to investigate other long-range projections in the auditory brainstem and beyond.

Channelrhodopsin-assisted circuit mapping (CRACM) is a precision technique for functional mapping of long-range neuronal projections between anatomically and/or genetically identified groups of neurons. Here, we describe how to utilize CRACM to map auditory brainstem connections, including the use of a red-shifted opsin, ChrimsonR.

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, ...

Neuroscience

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures the interaction of homologous chromosomes is the assembly of the synaptonemal complex (SC) in meiotic prophase I. Even though there is little sequence homology between protein components within the SC among different species, the general structure of the SC has been highly conserved during evolution. In electron micrographs, the SC appears as a tripartite, ladder-like structure composed of lateral elements or axes, transverse filaments, and a central element. 
However, precisely identifying the localization of individual components within the complex by electron microscopy to determine the molecular structure of the SC remains challenging. By contrast, fluorescence microscopy allows for the identification of individual protein components within the complex. However, since the SC is only ~100 nm wide, its substructure cannot be resolved by diffraction-limited conventional fluorescence microscopy. Thus, determining the molecular architecture of the SC requires super-resolution light microscopy techniques such as structured illumination microscopy (SIM), stimulated-emission depletion (STED) microscopy, or single-molecule localization microscopy (SMLM). 
To maintain the structure and interactions of individual components within the SC, it is important to observe the complex in an environment that is close to its native environment in the germ cells. Therefore, we demonstrate an immunohistochemistry and imaging protocol that enables the study of the substructure of the SC in intact, extruded Caenorhabditis elegans germline tissue with SMLM and STED microscopy. Directly fixing the tissue to the coverslip reduces the movement of the samples during imaging and minimizes aberrations in the sample to achieve the high resolution necessary to visualize the substructure of the SC in its biological context.

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex.

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures ...

High-Accuracy Correction of 3D Chromatic Shifts in the Age of Super-Resolution Biological Imaging Using Chromagnon

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