# 3. Tips for Data Collection¶

Data collection for use in 3D image processing softwares can be a little different than for other uses. In most cases, people optimize data collection so that individual slices look good to the human eye. This is not always the best for 3D data analysis and visualization. The following are some tips to help get the most out of the images:

• 16 bits images. If possible collect 16 bits per channel, instead of 8 bits. Although the pictures may look no different, the 16 bits images will have higher dynamic range, and it will be easier to extract features in darker areas of the image. If the microscope only supports 12 bits, that will still be better than 8 bits.
• More slices, less averaging. A common technique used to improve the image quality on confocal microscope is frame or line averaging. In this mode, the microscope takes each line/frame many times (typicalle 2 or 4 times), and creates an image by averaging each pass. Although this improves the image quality, it also increase the acquisition time, leading to problems such as: bleaching, tissue damage, movement during acquisition, … and to avoid these, a common technique is to reduce the number of slices taken. When acquiring data for 3D image processing software such as LithoGraphX, it is recommended to favour increasing the number of slices rather than averaging. The noise will be reduced later using neighboring slices.
• Dynamic range. When acquiring images, it is best to maximize their dynamic range: that is the difference between the brightness and the darkest pixel. To help you optimize this, most microscope’s software have a mode in which the saturated and black pixels are marked. When in that mode, first set the offset down until you see a lot of black pixels, and then increase it slowly to reduce their number until only a few are left. Then, increase the gain until many pixels are saturated, and decrease it slowly until only a few pixes are saturated. If you are interested in the geometry of cells, it is a good idea to saturate the cell walls slightly more: that will avoid issues with drops in signal and the exact shape will still be found at the middle of the saturated areas.
• Start and end slices shouldn’t contain any sign of your sample. It is tempting to start the acquisition when the sample is just visible, and stop just before it disappear. This is a bad idea: the reconstruction algorithms need the extra information that “there is no more tissue here” (or at least, not of interest) to reconstruct their geometry properly. So make sure you start a few micron before your sample, and stop a few micros after.
• Use Fiji to get TIFF files. Most microscope formats are proprietary. The Loci Bioformat group has reverse engineered the formats of the most popular microscopes and made plugins for ImageJ. We recommend Fiji, which is a distribution of ImageJ that contains these, along with many other useful plugins. Even if your microscope generates TIFF images, you might want to open them with the Bioformat plugins to extract the thickness of the slices. LithoGraphX only recognise ImageJ generated TIFF and OME TIFF for this.
• If voxel sizes are wrong. The TIFF format has no standard way to specify the voxel size in Z. ImageJ defines its own attributes for this, and the OME tif format another one. LithoGraphX recognises both formats. But some manufacturers provide their own TIFF file, with a different specification of the slice thickness. In this case, LithoGraphX will be unable to read it. You will need to use the Stack process Change Voxel Size in the Canvas folder and specify the z resolution.