|
|
This reference manual undergos revision as the program develops. A short form of the manual will appear as a chapter in the upcoming 3rd edition of The Proteomics Handbook, John Walker (ed) to be published in 2004 [Lemkin2004].
The following summaries list some information on using the interactive controls to run the program. You interact with the Flicker program using the mouse movments, keyboard commands, checkboxes states, slider controls parameter values, and pull-down menus. Figure 1 shows a screen view of the Flicker program.
If it appears that a spot and the surrounding region do match, then one has more confidence that the objects are the same. This putative visual identification is our definition of matching when doing a comparison. Full identification of protein spots requires further work such as cutting spots out of the gels and subjecting them to sequence analysis, amino-acid composition analysis, mass spectrometry, testing them with monoclonal antibodies, or other methods.
Using the mouse, the user initially selects (by clicking on the image - left or right window) what they suspect is the same prominent spot or object in similar morphologic regions in the two gel images. If you click on the spot with the Control key pressed, it will center that image at that spot in the window and also center it in the flicker window. You can continuously adjust the position in the flicker window, without changing the selected window position, by selecting the image you want to recenter and draging the mosue. When these two local regions come into alignment, they appear to pulse and the images appear to fuse together. At this point, differences are more apparent and it is fairly easy to see which spots or objects correspond, which are different, and how they differ. We have found that the user should be positioned fairly close to the flicker window on the screen to optimize this image-fusion effect (i.e., it does not work as well standing back more than a few feet from the screen).
Another useful trick is change the image canvas size after doing the initial alignment. This may help you focus on a smaller region after you have done the rough alignment. You can either use the "+" / "-" buttons under the parameter scrollbars (Figure 1.I), or you can use the Control-keypad + and Control-keypad - keys.
When comparing a light spot in one gel with the putative paired darker spot in the other gel one may want to linger longer on the lighter spot to make a more positive identification. Because of this, we give the user the ability to set the display times independently for the two images (typically in the range of 0.01 second to 1.0 second with a default of 0.30 second) using separate Delay scroll bars located under each image (Figure 1.E). If the regions are complex and have a lot of variation, longer display times may be useful for both images. Differential flicker delays with a longer delay for the light gel are useful for comparing light and dark sample gels. This lets you stare at the lighter spots to have more verification that they are actually there.
These flicker delay values are saved for the left and right images when you save the Flicker state.
Click to access DB
was enabled, it
will try to locate that spot in the associate Web server for that
image (see Figure 2).
a) 
b) 
Click to access DB checkbox. Then
set the (View | Use protein DB
browser, else lookup ID and name on active images) menu checkbox
command is enabled. Finally, click on the spot of interest and this
brings up the specific protein
annotation page from the active DB server (in this case
SWISS-2DPAGE) in a Web browser. Figure 3
shows an illustration of procedure.
In place of the Web browser, you can just have it report the
Swiss-Prot ID and the protein name annotation from the Swiss-2DPAGE
Web server. Set the (View | 
Use
protein DB browser, else lookup ID and name on active images) menu
checkbox command is disabled. Then when you click on a spot, it will
now try to get the Swiss-Prot ID and the protein name by the spot's
coordinates and display the results in your report window.
The next discussion describes looking up the (ID, protein names) for a set of spots you have defined.
Click to access DB
checkbox. Finally, click on the spot of interest and this brings up
the Web page from the active DB server (in this case SWISS-2DPAGE).
If the (View | Use protein DB
browser, else lookup ID and name on active images) is enabled,
then clicking on a spot will popup a Web browser with the associated
database page (as is shown). If the checkbox is off, then it will get
the data from that web page and report it. If you have defined a set
of spots in a spot list in an active gel, the (Quantify | Measure
by Circle | Lookup Protein IDs and Names from active map server
(selected image)) menu command will try to lookup each spot in the
list by its coordinates and will save the results (Swiss-Prot ID and
protein name) as the spot annotation's (id,name).
If you then have a set of corresponding spots you have defined in your gel, you can then edit their annotation so it matches the reference gel that you just assigned putative spot identifications. This is described in the following Figures 3.1a through 3.1d You can lookup the protein (ID,names) for a set of spots you have defined in the active gel map image that is link to federated 2D gel databases. See the vignettes for query a spot's putative identity and assigning a spot's putative identity for more details.
a) 
Use
'spot identifier ' for spot annotations) menu checkbox. Then
enable the Click to access DB
checkbox. Then, if you are connected to the Internet, with the active
gel selected, use the (Quantify | Measure by Circle | Lookup
Protein IDs and Names from active map server (selected image))
menu command. This will try to lookup each spot in the list by its
coordinates and will save the results (Swiss-Prot ID and protein name)
in each the spots annotation's (id,name).
b) 
Click to
access DB checkbox so that it does not popup the Web browser when
you then click on the image.
c) 
d) 
...) checkbox command. You may select
either SWISS-2DPAGE, UniProt, iProClass or iProLink servers - or none
of these. If you measure a spot (select a spot in an active image and
then type C-M) (and are connected to the Internet), it will
also lookup the Swiss-Prot protein (accession name, and protein id) on
the SWISS-2DPAGE server. Then, if you enable "Click to access DB", it
will pop up the particular active DB server you have selected. Figures
3.2a UniProt,
3.2b iProClass, and
3.2c iProLink.
An additional option that makes this easier to use is to enable the
(Edit | Auto measure, protein
lookup and Web page popup) checkbox command. Then when you click
on a spot in an active image (associated with a Web database), it
will: 1) measure the spot and add it to the spot list; 2) lookup the
Swiss-Prot (name, id); and 3) pop up the Web server on the currently
selected active DB server.
a) 
Use PIR UniProt DB access) checkbox
command set, enabling (Edit | Auto
measure, protein lookup and Web page popup) checkbox command, and
then clicking on the spot. b) 
Use PIR iProClass DB access) checkbox
command set, enabling (Edit | Auto
measure, protein lookup and Web page popup) checkbox command, and
then clicking on the spot. c) 
Use PIR iProLink DB access) checkbox
command set, enabling (Edit | Auto
measure, protein lookup and Web page popup) checkbox command, and
then clicking on the spot.
Flicker (C-F) checkbox enables/disables flickering.
Click to access DB checkbox enables/disables access to a
Web database server that is associated with a clickable image - if it
exists for the selected image. Turning on this option will disable
flickering.
Allow transforms checkbox enables/disables image transforms
Sequential transforms
checkbox enables/disables using the last image transform output as
input for the next image transform (image composition) if the Allow transforms is enabled
Some of these transforms involve spatial warping, which maps a local
region of one image onto the geometry of the local region of another
image while preserving its grayscale values. Another useful operation
is contrast enhancement that helps when comparing light or dark
regions by adjusting the dynamic range of image data to the dynamic
range of the computer display. Figure 2
shows the effects of contrast adjustment. Other transforms include
image sharpening and contrast enhancement. Image sharpening is
performed using edge enhancement techniques such as adding a
percentage of the gradient or Laplacian edge detection functions to
the original grayscale image. The gradient and Laplacian have higher
values at the edges of objects. In all cases, the transformed image
replaces the image previously displayed. Flicker will normally
transform the input imate to an output image. You can use another
transform on the previously transformed image if you had set the Sequential transforms checkbox.
Allow
transform enables/disables transforms, and Sequential transforms allows using
the previous transform as the input to the next transform, i.e., image
composition. The original image is denoted iImg. If you allow
transforms and are also composing image transforms, you may optionally
use the previous transformed output image (denoted oImg) as
input to the next image transform. The output (either iImg or
oImg) is then sent to the output1. Then the output2 which is
either zoomed or not is sent to the brightness-contrast filter if
active (specified by dragging the mouse in the selected window with
the SHIFT-key pressed). The output2 of the brightness-contrast
filter is denoted as bcImg. If you have never used the zoom or
brightness-contrast filtering since loading an image, then zImg
and bgImg are not generated and hence not used in the displayed
image. Figure 5. illustrates these four cases.
a) If no transforms or brightness-contrast filtering is used
on the selected image
(No transforms)
iImg -----------------------------------> output1
b) Image transforms may be composed from the original image (iImg) or
from Sequential composition of image transforms on the
selected image
(optional sequential)
+----------------------+
| |
V |
iImg -----> Transform -----------> oImg --> output1
c) The image may be optionally zoomed if the magnification is not 1.0X
on the selected image
(zoom)
output1 --------> zImg ------------> output2
or
(No zoom)
output1 ---------------------------> output2
d) Using brightness-contrast filter on output image
(BC filter)
output2 -----------------> bcImg --> display
or
(No BC filter)
output2 ---------------------------> display
Flicker (C-F))
a) 
b) 
c) 
Display gray values (C-G)) option. It
reports the total measurement circle density if the (Quantify |
Use sum density else mean
density)) option is enabled.
a)
b) 
c) 
Show report popup) command or the
Report scroller values button underneath the parameter
scrollers.
In both systems, you can specify additional command line arguments in Windows .bat and unix .sh scripts below.
REM File: Flicker-startup.bat REM Simple Flicker batch script for Windows. REM Starts Flicker from its jar file on the command line. REM To use more memory on startup, increase 96 (Mbytes) to a larger value. java -Xmx96M -jar Flicker.jarand
#!/bin/sh # File: Flicker-startup.sh # Simple Flicker batch script for Unix. # Starts Flicker from its jar file on the command line. # To use more memory on startup, increase 96 (Mbytes) to a larger value. java -Xmx96M -jar Flicker.jar
If you have your own gels (JPEG, GIF or TIFF formats), you can try loading them. You may want to limit resolution by first decreasing their size using an image editing program like Adobe PhotoShop or the shareware program ThumbsPlus (www.cerious.org ). Large very high resolution images that are 20Mb to 40Mb will not work as well. We suggest reducing the size to about 1Kx1K for good interactivity if you have any problems with running out of memory or very sluggish response. These image editing programs can also be used for converting other formats to JPEG, GIF or TIFF formats that Flicker can read.
An Internet connection is required to download the program from the Flicker Web site. New versions of the program and associated demo data will become available on this Web site and can be uploaded to your computer using the various (File | Update | ...) menu commands. If you have obtained the installer software that someone else downloaded and gave to you, then you do not need the Internet connection to install the program. If you will be using the active gel image maps associated with federated 2D-gel databases, then you will need the Internet connection for accessing those databases. You do not need the Internet for local image comparisons.
If you are using the Windows Flicker.exe file to start the program or clicking on the Flicker.jar file, you can't change the default startup memory.
If you are using the Flicker.jar in a script using the java interpreter as in the following example which uses the -Xmx96M (specifying using 96 at startup). Change 96 to a larger size if you want to increase startup memory.
java -Xmx96M -jar Flicker.jar {additional command line args}
For example, if you have four images image1, image2, image3 and image4 in your directory MyData-folder, then the submenu entries will be
Note that it does not generate comparisons between directories. You can still do that by clicking on the left (and then later the right) image and using the (File | Open image file) command to manually load the file into the image. If you want to do this often, put your images in the same directory.
If you have this type of data, it will also add the (File | Open user images | List user's images by directory) command. You can use this to get a report of all of the files in the popup report window.
You will not use the (Files | Update | Flicker program) command. Using this command will get the latest release which is not what you want if you want an earlier release.
Instead do the following. You copy the version of Flicker.jar you want to the location on your disk where you had installed Flicker overwriting the previous version. For example, on MS Windows this is typically
C:\Program Files\Flicker\Flicker.jar
If you installed Flicker elsewhere on your computer, then that is
where you need to change it. Go to the files distribution in the File
mirror web page.There are a number of .jar files of the form
Flicker.jar-V0.xxCopy the one you want to your computer.
To be used with the Flicker startup process, this file must be called Flicker.jar, so you must rename it as Flicker.jar. The procedure is as follows:
DB/ and Images/The DB/ folder must have two files FlkDemoDB.dir (the list image images in the Images/ folder, and FlkDemoDB.txt which is a tab-delimited spread sheet describing the demo images. NOTE: demo images generally live in the top level of the Images/ directory. They are not in subdirectories of the Images/ directory. If you are designing such a remote demo Web site, you could model the files from the current Flicker DB files.
Once you are sure your remote web site is set up correctly, then simply use the (File | Update | Add user's Flicker Demo Images DB by URL) command to download and and update the Flicker demo database. After it is successful, it will tell you to exit and restart Flicker. If you go to the Files | Open demo images| ... menu, you should see the new demo data you have installed.
The Flicker.properties file is the user-specific preferences for generic properties such as colors, view options, etc. This file is created when you exit flicker and read (if it exists) when you start it. If you have never run flicker or the file was deleted, Flicker uses the default values. If the Flicker.properties file exists, it overides these files. If you have started Flicker using a .flk startup file, this in turn overides any settings from the Flicker.properties file. For example, you may want to use a red trial object cursor instead of yellow, etc. If you have both a Flicker.properties file and are starting Flicker using a .flk startup file, it (1) reads the user preferences first from Flicker.properties, and then (2) reads the .flk startup file to overide these preferences. If you want to reset the standard view, use the (Edit | Reset default view) command to overide the previous view read from Flicker.properties. If the you ever have problems starting Flicker because of a corrupted Flicker.properties file, just delete it (it is kept in the same directory where you installed Flicker) and restart Flicker. When you exit, it will create a new properties file.
'. Selections prefaced with a '' and indicate '' indicate that the command is a checkbox
that is enabled and disabled respectively. Selections prefaced with a
'
' and indicate '
' indicate that the command is a
multiple choice "radio button" that is enabled and disabled
respectively, and that only one member of the group is allowed to be
on at a time. The default values set for an initial database are shown
in the menus. Selections that are not currently available will be
grayed out in the menus of the running program. The command short-cut
notation C-key means to hold the Control key and then
press the specified key.
- load pairs of
demonstration gel images
- load single
(selected image) or pairs of user's image data if they had set
up the Images/ directory as
specified on their data
- load active gel
image from the Internet in selected image
- update Flicker
programs and data from open2dprot.sourceforge.net/Flicker
server
- change the size
of the 3 canvases and the overall Flicker window
- change overlay
graphics colors
-
to change the target color
-
to change the trial object color
-
to change the landmarks color
-
to change the circle mask measurement color
-
to change the image guard region color
Use linear else log of TIFF files > 8-bits - transform
the input TIFF image if it is more than 8-bits of intensity.
Enable saving transformed images when do a 'Save(As)
state' - If you have transformed the left and/or the right
image(s) and are allowing transforms, then this will save the
transformed images in a tmp/ directory as .gif files
when you save the Flicker state.
Use protein DB browser, else lookup ID and name on active
images - used when click on a spot in an active DB that has
click to access DB
checkbox enabled. If this command is set, report the protein
identication in a popup window. If it is not set, then lookup
the protein ID and name and report it in the popup Report window.
Auto measure, protein
lookup and Web page popup - if one the following active DB
servers is enabled, it will: 1. measure the spot and add it to
the spot list; 2. lookup the Swiss-Prot (name, id); and 3. pop
up the Web server on the currently selected active DB server.
- SWISS-2DPAGE, UniProt, iProClass or
iProLink servers. If you measure a spot (select a spot in an
active image and then type C-M) (and are connected to
the Internet), it will lookup the Swiss-Prot protein (accession
name, and protein id) on the Swiss-2DPAGE server. Then, if you
enable "Click to access DB", it will pop up the particular
active DB server you have selected.
Flicker images (C-F) - enables/disables flickering.
-
enable/disable various overlays for the images.
View landmarks - add landmarks to the overlay display in
images
View target - add target to the overlay display images
View trial object - add trial object to the overlay display
images
View boundary - add boundary to the overlay display images
View Region Of Interest (ROI) - add ROI to the overlay
display images
- set various measurement overlay
options. Some of these require the (Quantify | Measure by
circle | List-of-spots else
trial-spot measurement-mode (C-J)) checkbox command be set.
View measurement circle - add measurement ("+M") and
background circles ("+B") to the overlay display images. The
("+M") is replaced with a series of spot measurements. These
are visible if the list-of-spots measurement mode is set.
Use 'circle' for measured spot locations - if you have
measured spots, show the location with a circle. These are
visible if the list-of-spots measurement mode is set. This
will show the actual circle masks (of different sizes if you
changed the sizes) around each measured spot. Otherwise, it
shows it as a '+' marker.
Use '+ for measured spot locations - if you have
measured spots, show the location with a '+'. These are visible
if the list-of-spots measurement mode is set.
Use 'spot number' for measured spot annotation - if you
have measured spots, add the spot measurement number
annotation. These are visible if the list-of-spots measurement
mode is set.
Use 'spot identifier for measured spot annotation - if
you have measured spots, add the spot identifier (if it exists
else <none>) annotation. These are visible if the
list-of-spots measurement mode is set. You may edit the spot
identifier using the (Quantify | Measure by circle | Edit
selected spot from spot list (C-E)).
-
enable/disable various multiple "ganged" image options where
performing a change will do the same change in several images
simultaneously. This is most useful after doing an initial
alignment, zoom, etc.
Multiple popups - make multiple popup windows instead of reusing
one window
Gang scroll images - move left and right images scrolling
together
Gang zoom images - zoom left and right images together. This
is useful if they are the same exact magnification
Add guard region to edges of
images This is useful for aligning spots that are along the
edge of the images while flickering. The guard region's color
can be changed via the Edit menu (Edit | Set colors | Guard
region color).
Display gray values (C-G) - show pixel gray values of cursor
trial object in the left and right image titles as well as (x,y)
coordinates. Also display the measurement circle if it is > 1 as
you move the mouse.
Show report popup - display the report popup window again if
needed
Use threshold inside [T1:T2] filter - filter by pixels inside
the range [T1:T2], otherwise pixels outside of [T1:T2]
Allow transforms
checkbox enabled.
Sequential
transforms checkbox disabled.
a) 
b) 
c) 
In the mean time, we refer to "density" as either of these two measurements. You may measure the total integrated density (summing the pixel gray values or OD), or mean density (the total density/#pixels in the region). This background density is used to correct the measurement density if it was defined (otherwise 0.0 is assumed). Note that you may clear the background measurement circle using the (C-W) command.
You can save the spot list in a tab-delimited file suitable for import to Excel, by 1) clearing the Popup Report Window; 2) select the image you want and use (Quantify | Measure by circle | List spots in the spot list (tab-delimited)) command; 3) then, Save the text in the Popup Report Window or cut and paste it into Excel, etc.
It is possible to generate a list of paired spots between the two images using the (Quantify | Measure by Circle | List paired 'id' annotated-spots in both spot lists (tab-delimited)) command. It lists paired spots that occur in the spot lists of both gels and have the same annotation 'id' values (case-sensitive). The user assigns the spot 'id' annotations using the (C-I) spot editing command to assign the case-sensitive spot identifiers. (C-E) command may also be used to edit the ids'. The paired spot data list may be exported to Excel similar to what was described in the last paragraph for spot lists.
Note that the background density is not the same everywhere on the gel image. Therefore if you are measuring spots in regions of the gel with quite different background (e.g., tail streaks from other spots, etc.), to get better estimated quantification you need to redefine the background adjacent to where you are making measurements. Recent versions of Flicker track the most recent background (C-B) estimate and associate it with new measured spots (C-M) until you redefine it.
- measure
intensity/density within the circlular mask
Use measurement counter) is
enabled. An alternative way to measure spots is to hold the
ALT-key when you press the mouse to select the spot. This
combines spot selection and measurement in one operation. Note
that you can toggle between trial- mode and spot-list measurement
mode using the (Quantify |
List-of-spots else trial-spot measurement-mode (C-J)) checkbox
command.
- measure
intensity/density within user-drawn boundary
- Print
data-window
20x20 - window size
Decimal - data format radix
- calibrate
optical density or other step wedge
Use demo leukemia gels ND wedge
calibration preloads wedge - used to demonstrate the ND step
wedge calibration process. We have built in four sample ROI's and
the corresponding OD values corresponding to the step wedge scanned
with the four demo gels (Leukemia-AML, Leukemia-ALL, Leukemia-CLL,
Leukemia-HCL). Enable this option before invoking the
Optical density by step wedge command.
-
region of interest operations
List-of-spots else trial-spot
measurement-mode (C-J) to either measure spots or use a trial
measurement mode. In measurement mode, add spot label overlays
consisting of measurement locations (either "+" or circles or
neither) followed by measurement annotation (measurement # and/or
spot specific identifier (if edited), or neither). As specified by
the (View | Set view measurement options | ... checkbox
options. In measurement mode it will number the measurements 1, 2,
3, ... etc, whereas in trial-measurement mode it will show a
single "+M" instead. Note that in trial mode, it will redraw the
measurement circle as you change the size with the slider. Note
that if you are measuring several different ROI's with
(C-R), then it will assign sequential measurment numbers to
these ROI measurements.
Figures 12a-12h show various permuations of the spot list overlay options. Figure 13 shows an example of a histogram of the Region Of Interest (ROI). Figure 14 shows examples of calibrating gray scale a step wedge scanned with the image using the step wedge calibration wizard. Figure 15 shows calibrating grayscale where there are calibration spots in the image by using the spot list calibration wizard.
a.1) a.2 b.1) b.2) c) d) e) f) g) h)
Panels (a.1) and (a.2) show a single spot measurement
image and report and does not create a list of measured spots. The
(Quantify | List-of-spots else
trial-spot measurement-mode (C-J)) should be turned off. The
VM: View measurement circle
option should be enabled.
List-of-spots else
trial-spot measurement-mode (C-J)) should be turned on. The VM:
Use 'circle' for measured spot
locations option should be turned on, VM: Use 'spot number, for spot annotations
option should be turned on.
List-of-spots else trial-spot
measurement-mode (C-J)) should be turned on. The VM: Use '+ for measured spot locations
option should be turned on, VM: Use
'spot number, for spot annotations option should be turned on.
List-of-spots else trial-spot
measurement-mode (C-J)) should be turned on. The VM: Use 'circle' for measured spot
locations option should be turned on, VM: Use 'spot number, for spot annotations
option should be turned on, and VM: Use 'spot identifier, for spot
annotations option should be turned on. The user had edited the
spot identifiers by (1) clicking on a spot, (2) doing the (Quantify
| Measure by circle | Edit selected spot from spot list (C-E))
command, and 3) typing a spot "identifier" for that spot and pressing
"Done". Spots where the 'id was not edited are indicated by
"<none>". For this example, we set 'id' to "SpotOfInterest" for
spot #1 and a space for spot #3. Spaces make the identifier disappear.
List-of-spots else trial-spot
measurement-mode (C-J)) should be turned on. The VM: Use '+' for measured spot locations
option should be turned on, VM: Use
'spot number, for spot annotations option should be turned on, and
VM: Use 'spot identifier, for spot
annotations option should be turned on.
List-of-spots else trial-spot
measurement-mode (C-J)) should be turned on. The VM: Use 'circle' for measured spot
locations option should be turned on, and VM: Use 'spot identifier, for spot
annotations option should be turned on.
List-of-spots else trial-spot
measurement-mode (C-J)) should be turned on. The VM: Use '+' for measured spot locations
option should be turned on, and VM: Use 'spot identifier, for spot
annotations option should be turned on.
List-of-spots else trial-spot
measurement-mode (C-J)) checkbox option should be turned on. The
VM: Use 'spot identifier, for spot
annotations option should be turned on.
Once an image is calibrated and the calibration saved (in the {installation directory}/cal/{imageFile}.cal file), it may be reused everytime the image is loaded into Flicker. The initial calibration is created from the step wedge data and known corresponding OD values using a histogram specified by a region of interest (ROI) specified as the step wedge region.
To make it easier to demonstrate this process, we have set up special demo calibration data for the four demo Leukemia gels. If you have enabled (Quantify | Calibration | Use demo leukemia gels ND wedge calibration preloads) checkbox, it will preload the OD values and step-wedge ROI for demonstration purposes. If you are using any other data, you must specify the OD values and step-wedge ROI. You invoke the wizard by the (Quantify | Calibration | Optical density by step wedge) command. This will popup the calibration wizard window.
a) 
b) 
c) 
a) 
b) 
View measurement circle) option. The
numbered image is shown in Figure 16a.You may also list the saved spots in the report window (shown in Figure 16b). Hint, press the Clear button in the report window first. Note that if you save the flicker state, then the spot lists will be saved in the .flk file state and associated .spt spot list files in the spt/ directory. Flicker will restore the spot lists if you start flicker on the saved .flk startup file.
List-of-spots else trial-spot
measurement-mode (C-J)) menu checkbox.
a) 
b) 
a) 
b) 
c) 
Flicker can then use the annotated spot lists you have generated to create a paired spot table shown below.
a.1) 
a.2) 
b.1) 
b.2) 
Windmp [183:203, 168:188] Center (193,178) 20X20 size,
sampled: 1 pixels, data-radix: Decimal
Image: Images\plasmaH.gif
X 183 184 185 186 187 188 189 190 191 192 193 194 195 196 197 198 199 200 201 202 203
Y --- --- --- --- --- --- --- --- --- --- --- --- --- --- --- --- --- --- --- --- ---
168 71 61 57 55 54 54 56 57 56 50 46 42 42 43 45 42 43 48 53 67 80
169 71 53 50 48 45 48 50 55 60 56 46 42 40 43 42 40 37 41 47 54 65
170 88 49 44 42 40 42 45 53 56 61 50 41 40 39 39 37 34 37 38 45 54
171 97 49 41 39 39 40 43 53 60 61 54 43 39 38 37 36 34 36 37 43 50
172 106 47 39 37 36 37 39 53 72 78 72 50 43 38 37 35 33 34 35 39 44
173 84 41 38 37 34 33 35 50 82 94 84 58 48 41 40 37 34 35 35 39 44
174 66 39 38 37 35 33 35 50 84 96 84 59 48 41 40 39 35 37 37 39 44
175 47 41 43 40 36 34 34 49 78 84 71 56 50 43 44 43 38 37 37 47 46
176 50 46 46 46 39 34 35 56 91 95 77 61 58 54 57 54 47 43 41 43 47
177 53 49 51 50 42 37 38 69 117 122 98 74 66 62 65 62 51 47 44 45 50
178 58 59 65 65 53 43 46 106 171 174 139 94 78 71 67 65 57 54 50 48 54
179 63 77 97 97 82 59 55 104 163 164 120 76 64 58 62 62 61 58 55 52 56
180 67 90 122 124 106 72 60 86 132 132 95 64 59 55 64 62 61 59 57 53 57
181 76 140 189 193 174 128 94 59 70 68 60 55 55 54 59 63 60 60 58 55 58
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this reference manual
As part of the demonstrations, we present a series of as short vignettes that have answers to specific "How do I do ...?" types of questions. There are several demonstration images that are available when you download and install Flicker. Look in the (File | Open demo images) menu. These commands will load pairs of demonstration images.
[2] Hood L. (2002) A Personal View of Molecular Technology and How It Has Changed Biology. J Proteome Biology 1, 399-409.
[3] Herbert B.R., Pedersen S.K., Harry J.L., Sebastian L., Grinyer J., Traini M.D., McCarthy J.T., Wilkins M.R., Gooley A.A., Righetti P.G., Packer N.H., Williams K.K. (2003) Mastering Proteome Complexity Using Two-Dimensional Gel Electrophoresis. PharmaGenomics 3, 21-36.
[4] Pedersen S.K., Harry J.L., Sebastian L., Baker J., Traini M.D., McCarthy J.T., Manoharan A., Wilkins M.R., Gooley A.A., Righetti P.G., Packer N.H., Williams K.L., Herbert B.R. (2003) Unseen Proteome: Mining Below the Tip of the Iceberg To Find Low Abundance and Membrane Proteins. J Proteome Biology 2, 303-311.
[5] Pieper R., Su Q., Gatlin C.L., Huang S.-T., Anderson N.L., Steiner S. (2003) Multi-component immunoaffinity subtraction chromatography: An innovative step towards a comprehensive survey of the human plasma proteome. Proteomics 3, 422-432.
[6] Pieper R., Gatlin C.L., Makusky A.J., Russo P.S., Miller S.S., Su Q., McGrath A.M., Estock M.A., Parmar P.P., Zhao M., Huang S.-T., Zhou J., Wang F., Esquer-Blasco R., Anderson N.L., Taylor J., Steiner S. (2003) The human serum proteome: Display of nearly 3700 chromatographically separated protein spots on two-dimensional electrophoresis gels and identification of 325 distinct proteins. Proteomics 3, 1345-1364.
[7] Anderson N.L., Anderson N.G. (2002). J Mol. Cellular Proteomics 1, 845-867; and J Mol. Cellular Proteomics 2, 50.
[8] Liotta L.A., Espina V., Mehta A.I., Calvert V., Rosenblatt K., Geho D., Munson P.J., Young L., Wulfkuhle J., Petricoin E.F. 3rd. (2003) Protein microarrays: meeting analytical challenges for clinical applications. Cancer Cell 3, 317-325.
[9] Hoving S,. Gerrits B., Voshol H., Muller D., Roberts R.C., van Oostrum J. (2002) Preparative two-dimensional gel electrophoresis at alkaline pH using narrow range immobilized pH gradients. Proteomics 2, 127-134.
[10] Von Eggeling F., Gawriljuk A., Fiedler W., Ernst G., Claussen U., Klose J., Romer I. (2001) Fluorescent dual colour 2D-protein gel electrophoresis for rapid detection of differences in protein pattern with standard image analysis software. Int J Mol Med. 8, 373-377.
[11] Lopez M.F., Mikulskis A., Golenko E., Herick K., Spibey C.A., Taylor I., Bobrow M., Jackson P. (2003) High-content proteomics: Fluorescence multiplexing using an integrated, high-sensitivity, multi-wavelength charge-coupled device imaging system. Proteomics 3, 1109-1116.
[12] Sanchez J.-C., Appel R.D., Golaz, O., Pasquali C., Rivier F., Bairoch A., Hochstrasser, D.F. (1995) Inside SWISS-2DPAGE database. Electrophoresis 16, 1131-1151.
[13] Appel R.D., Bairoch A., Sanchez J-C., Vargas J.R., Golaz O., Pasquali C., Hochstrasser D.F. (1996). Federated two-dimensional electrophoresis database: A simple means of publishing two-dimensional electrophoresis data. Electrophoresis 17, 540-546.
[14] Appel R.D., Sanchez J-C., Bairoch A., Golaz O., Miu M., Vargas J.R., Hochstrasser D.F. (1993) SWISS-2DPAGE: A database of two-dimensional gel electrophoresis images. Electrophoresis 14, 1232-1238.
[15] Appel R.D., Sanchez J.-C., Bairoch A., Golaz O., Rivier F., Pasquali C., Hughes G.J., Hochstrasser, D.F. (1994) SWISS-2DPAGE database of two-dimensional polyacrylamide gel electrophoresis. Nucleic Acids Res. 22(17), 3581-3582.
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Revised: 05/12/2007 |