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    • CommentAuthorredsox07
    • CommentTimeAug 5th 2007 edited
     

    This protocol explains how to: use Image-Pro Plus to measure neurons in the in vitro TTX experiment.

    Protocol revised: 8/5/2007

    Protocol written by: Mike Neri

    Measuring Neurons from the in vitro experiment with Image-Pro Plus

    *Only the 4 computers down the center aisle of Dana 220 have Image-Pro Plus installed on them (unless you remove the yellow key and use in on one of the computers in the Lom Lab) so you must first log on to one of these computers to be able to use this program.

    1) Go to the Start Menu, select “Image Pro Plus 5.1”

    2) In the “Select Menu and Toolbar” box that comes up, select “Complete” and press “OK”

    3) Click the “X” in the top right corner of the Macro Player box that pops up to close out of that box

    Calibrating the program for measuring neurons (this only needs to be done once when you open the program at a computer, unless it doesn’t come up the next time you open Image-Pro):

    *You must first have taken a picture of the scale bar on the phase setting of the fluorescent microscope using the correct objective (40X) and saved it as a .tif file

    1) Go to “File” -> “Open” and open the .tif picture file that you took of the scale bar

    2) Go to “Measure” -> “Calibration” -> “Spatial Calibration Wizard”

    3) In the “Spatial Calibration Wizard” Box that pops up, select the “Calibrate the Active Image” under the “I would like to:” prompt and select “Next >”

    4) Create a name for your calibration, set the units to microns (ums), and put a check in the “Create a Reference Calibration” box and the “Set as System Calibration” box (which will pop up when you check the first box) and click “Next >”

    5) In the next window, click on the “Draw Reference Line” button

    6) A green line will pop up on your picture as well as a “Scaling” box.  In the scaling box, make sure the number under the “Represents how many units?” question is 200.  Then draw a reference line as closely as you can to cover 20 units on the scale bar.

    7) You can zoom in and out using the scroll button on your mouse to get it more exact.  Once you have a line drawn that you are satisfied with, click “OK” in the scaling box.

    8) Under the “Draw Reference Line” button, take a look at the “Calibration Report.”  This tells you how many microns will be represented per pixel.  I always made sure that my calibrations were the same value, and that was 0.261053 um/pixel.  You may want to try for this number, or at least keep it constant throughout your data as well as it will allow for more consistent results.

    9) Once you are satisfied with you calibration, click “Finish” (you can redraw your calibration line if you are not happy with how it turned out).  You can now close out of your calibration picture and you are ready to measure neurons.

    Important Note: You must click “Apply” in the “System Spatial Calibration” box that is now on your screen before you export any of you data to excel.  You must do this for each neuron or you will not have the correct length measurements.  If you forget to do this before you start measuring your neuron, you can click “Apply” and then click the “Update” button in the “Measurements” box to make sure all of your measurements are calibrated.  If you have no idea what I am talking about right now, keep reading and you will understand.

    Once your program is calibrated, you are ready to measure your neurites.

    1) At the top of your screen, go to “Sequence” -> “Merge Files”

    2) In the “Open File” Box that pops up, select all the built images from a single neuron and click “Open.”  This will create a movie of sorts composed of multiple frames with different views of a single neuron so you can benefit from all the different pictures and builds of a single neuron.

    3) A screen with your pictures has popped up, along with the “Sequence Toolbar.”  You can cycle through your pictures using the button that looks like an arrowhead pointing at a line and choose the view at which you can best see the neurites at that point.  You can also zoom in on the neurites using the roller ball on your mouse if you have selected the picture window at the time (the roller ball will not move you up and down in the picture like on a web page, so you must grab and move the side bar to do so).  I like to scroll in and then increase the size of the picture window to take up most of the view.

    4) Now at the top of the window select “Measure” -> “Measurements.”  The “Measurements” box will pop up on your screen.  Make sure the “Features” tab is selected near the top of that box and choose the squiggly line from the “Features” section on the left side.  It is the 3rd choice down in the 3rd column (the right column).

    5) There may be instructions that pop up about how to use the squiggly line tool, which you can read and then click out of.  I then use the “Sequence Toolbar” to select the green fluorescence picture to start and zoom in pretty close to the cell body.  Then using my mouse I make one left click at the point where it appears to me that the cell body ends and the primary neurite begins.

    6) A gray line will appear from the dot where you clicked.  You can then drag your mouse around and left click again at the next spot along the neurite so that the line will accurately outline the length of the neurite.  Once a line is set in place, it will turn red and allow you to move the mouse out from that point.  Keep moving along the neurite until you reach the end.  I find the phase view the most useful as I move away from the cell body further along the neurite.

    7) When you come to a fork in the neurite, choose to follow the path of the neurite that seems most logical based on the width, direction, etc. that the neurite is traveling.  I didn’t try to just follow the longest neurite from a branch point because that was not always the one that seemed the most logical extension of the path of the one I was following.  Also, when you are clicking along a neurite, be careful not to double click, as that will cause the program to try to outline whatever object you are on.  If you do so by accident, select the regular arrow tool in the “Feature” section of the “Measurements” box (which should cause your line to be erased) and then select the squiggly line again and start over (I have not yet discovered an “undo” function in this program, which can make things frustrating).

    8) When you reach the end of a neurite, and have all the length outlined, I like to cycle through all the views of the neurite using the “Sequence Toolbar” to make sure I am truly at the end.  Oftentimes a neurite will be easier to view at a certain part in one picture than it is in another, so you may find it helpful to cycle through different pictures as you are outlining the neurite as well.  Once you are convinced that you have reached the end, pause a few seconds (otherwise the program often will erase the last segment of the line that you drew) and then click the right mouse button one time to complete your line (in this program, each line that you draw is referred to as a “feature” and shown in the “Features” tab of the “Measurements” box).

    9) If there is a growth cone at the end of your neurite, I draw the line out to the center of the growth cone and end the neurite there.

    10) If you are not satisfied with this line, you can select it using the regular mouse button found in the “Features” section of the “Measurements” box and then press “Delete” on the keyboard or click the black “X” in the “Features” section of the “Measurements” box.

    11) After the primary neurite is completely outlined, I work my way back toward the cell body and outline any secondary and tertiary neurites that branch off of it.  (Secondary neurites are neurites that branch off of primary neurites (ones that come directly from the cell body), tertiary neurites branch off of secondary neurites, etc.)  As I am working my way back, I also cycle through the different pictures using the “Sequence Toolbar” to make sure I am not missing anything as I go.

    12) You can also improve you ability to see the neurites for measurement by opening the “Contrast Enhancement” box by clicking on the icon along the top of the program that looks like three little lines with bars at different levels across them (the one with bars at the same levels will reset the contrast to the original levels when the files were opened).  You can then move the bars up and down to improve your ability to see the neurites.  This is why you don’t want to mess with the contrast in Photoshop when you are building neurons: it will limit your ability to alter the contrast here.  If you don’t like how it is altered, you can start over by clicking the “Reset” button in the pop up box or the button with the bars all at the same level in the top toolbar.

    13) Continue to measure the neurites until all are covered in a line.  Do not ignore the really short neurites that you see coming off of a neurite – these are often spikes, which are neurites that are less than 5um in length and which we count.  However, not everything that appears to come off of a neurite is a branch or a spike, so the measurer needs to decide which is which and be consistent.

    14) Once all neurites, including spikes, are measured using the squiggly line tool, I like to make sure that my calibration is applied by clicking “Apply” in the “System Spatial Calibration” box and then clicking “Update” in the “Measurements” box.  This just makes sure that all of my lengths are going to be calibrated correctly.

    15) Open Microsoft Excel and create a file for each separate experiment.  Name it something like “24hrs. TTX Expt (insert experiment number) Data.xls.”  Next, open up the Microsoft Word template file that I created, which can be found by following this path: P:\Biology\Lom\misc\biologylom\TTX24hr.  The file name is something like “In vitro Data Sheet Template (go Red Sox).doc,” but may have been altered by the Yankee-loving Barbara O to say something about the Yankees in the parentheses.  These two files are where you can collect and organize your data.

    16) Select the “Input/Output” tab in the “Measurements” box.  In the “Export Data” section, make sure the black dot is in the circle next to “Feature” under the “Data to Output:” section and “DDE to Excel” under the “Output Data to:” section.  This will output a whole bunch of data to your open excel file, which will always appear at the top of the sheet, so you will want to leave about 40 rows blank at the top of you excel sheet for data to be imported every time.  Click the “Export” button.

    17) The good news is that of all the data that you just saw appear in you excel file, you only need the first column, the “Features” column, and the “Length” column (should be columns A, B, and F).  Delete all the other data, then move the “Length” data into column C next to the “Features” column (highlight the “Length” column, press Ctrl+X, select the top cell in column C, then press Ctrl+V).  This will put all the data you need in a nice, compact three column set.

    18) Move all that data to the appropriate part of the data sheet (remember, leave at least 40 rows blank at the top of your excel file).  Label the data in the blank top cell of the first column with the neuron name.  In the first cell underneath the last number in the first column, write “Total length,” under that write “# tips,” under that write “# primary,” and under that write “# spikes.”  These are the 4 things that you are going to keep track of.

    19) To tabulate the total length, highlight all the lengths from the length column and press the “AutoSum” button at the top of your excel file (it looks like the Greek letter Sigma (Σ)).  This should add all the lengths together and put the total length in the space across from where you typed total length (for the first time, I would add the lengths on a calculator on in your head to make sure the button is functioning correctly, but after that you can trust the program). 

    20) For the number of branch tips, it should just be the bottom number in the first column (which just shows the number of features imported).  It is not necessarily the same as the number next to the “T” in the “Feature” column.  If you had to delete a feature because it was drawn incorrectly, it will just skip that feature number and give you and the next number number.

    21) To tabulate the number of primary branches, just go back and look at the picture in Image-Pro and count how many branches come out of the cell body.

    22) To tabulate the number of spikes, look in the length column and count how many have lengths of less than 5 um and record it in the appropriate place in your excel file.

    23) Finally, in the first column, record what type of branch each feature is.  You can do this by looking back at the Image-Pro window and seeing whether a branch is a primary, secondary, tertiary, quaternary, etc.  Then, where the branch number currently is (note: not the feature number that has the “T” before it, but the branch number in the first column) write “1” or the phrase “1 branch” (standing for primary branch) for all the primary branches.  Do the same for the other branches, using “2,” “3,” “4,” to mean secondary, tertiary, quaternary, etc.  This will give you an idea of the complexity of the neuron.

    24) Now, here is how to backup your data and pictures into the convenient Word file template I have created.  Highlight your total length and press Ctrl+C to copy it, go to the data sheet template and paste it into the total length space (be sure to also put the correct neuron name at the top of each text box).  Then just type the other, less complicated numbers into their respective places in the document.

    25) The space below the data you just imported is for you to paste a picture of your neuron for future reference.  When you save your movie in Image-Pro, it does not save the lines that you have drawn on it, so you cannot compare which feature corresponds to which line.  Therefore, we paste a picture from Image-Pro into this document and we can save it.  So go to Image-Pro, make your neuron picture as big as you can, use the green fluorescent picture as the backdrop (choose using the Sequence Toolbar – it doesn’t have to be the green picture, but I found it most useful because it provided good contrast for the yellow lines to show up on), and press and hold the “Shift” key and then press and release the “Print Screen” key.  Then go back to you Word document, paste the picture into the appropriate text box, click on the picture, and using the crop tool (can be found from the picture toolbar, which can be found by going to “View” -> “Toolbars” -> “Picture”) begin in the corner and crop the image so that only the neuron is visible.  You can then resize the image to that it fits into the whole text box.

    26) At this point, you want to save the movie in Image-Pro in a file for future reference (even though the yellow lines will be lost).  I saved mine here: P:\Biology\Lom\misc\biologylom\TTX24hr Expts\24 hr Expts ANALYSIS using files names like this: “Expt 1 Built Movies.”  The files names had the neuron name and then “builtmovie” at the end, like this: “1A01builtmovie.tif.”  This is also the folder where I saved my excel and word files (as well as backing them up onto the desktop of the computer I was working on).

    27) At this point, I record all the data in my notebook, putting the neuron name, total length, number of branch tips, and number of spikes as my final way of backing up the data.

    28) Now you are ready to start on your next neuron by opening it up from “Sequence” -> “Merge Files” etc.  It seems like a lot of busy work saving the data this way, but you will be thankful at the end when your data is neat, organized, and complete.  Please update and change this protocol in any way or if you discover any ways to fix the little problems that I noted that I encounter along the way.

    Good luck with you neuron measurements!

    Mike, summer 2007

  1.  
    When everyone first starts measuring, individual differences in technique can translate into significant variations in final measurements. The number of spines recorded and the starting/end points for each are some of the possible sources of variations. If your group decides you need to compare decisions made while measuring, it might help to simply highlight the small spikes you recognized during measurement (using something like a transparent circle over each spike using Word drawing tools). If you label your spikes (in your word document images) and make a note of which measurement they each correspond to, you'll be able to compare everyone's spike recognition and measurement really easily and quickly. You won't have to go back and waste any time trying to remember which measurement belongs to which spine and so on.

    Also, if you are having a hard time finding spines or the end of neurites in the fluorescent image, be sure you are playing with contrast, brightness, etc... in the green channel. You can select which channel you are manipulating by choosing one of the colored buttons immediately below the contrast, brightness and gamma toggle bars (Enhance Menu --> Contrast Enhancement).
    • CommentAuthordoippolito
    • CommentTimeDec 20th 2007
     
    When measuring numerous neurons back to back, you generate a lot of data and I found that using the DDE options to export to Excel was a bid tedious. I saved a lot of time by exporting to the "Clipboard" instead of the DDE route and just pasted the entry into the next available cell in the spreadsheet (if you are unfamiliar with the clipboard, exporting data there is equivalent to copying('control+c'-ing) the measurement data. All you have to do is paste the data into excel like you would a normal selection).
    • CommentAuthorjuruble
    • CommentTimeJan 4th 2008
     

    To simplify the "Export" of your data to Excel from ImagePRO further, change #16 in the protocol to the following.  I find this simpler than copying the data to the clipboard and pasting, since it cuts those two steps down to one.

     

    16) Select the “Input/Output” tab in the “Measurements” box.  In the “Export Data” section, make sure the black dot is in the circle next to “Feature” under the “Data to Output:” section and “DDE to Excel” under the “Output Data to:” section. 

    Then select "DDE Options."  This window will allow you to specify for ImagePRO to paste each new data set directly below the last one, so you don't have to keep 40 spaces open at the top of your Excel file (as prescribed in the original protocol), which can be tedious.  Click the button next to "Increment position for next data set by:" and enter "30" in the rows column, which tells ImagePRO to put each new data set 30 rows below the previous data set in Excel.  Click the green "check" mark to save this. 

    THEN specify the row you want to start at (this is the row where ImagePRO will paste the first data set you export) by typing in the number of the row below "Position data set."  For instance, if you are just starting your Excel file, the row will be set at "0," but if you are continuing an existing Excel file and the last data set ends at row 200, you can enter something like "230" so that the first data set you export will be well below the previous data in the file.  YOU MUST DO THIS AFTER YOU SET THE # OF ROWS TO INCREMENT DATA, above.  Otherwise it will just change back.  Click OK.


    Now you can click the “Export” button to export your data to Excel.  You don't need to set DDE Options every time -- just for the first data export of each session.  Then you can just click "Export" each time and it will automatically increment the data set below the previous one by the number of rows you specified.