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Lecture

BIOB12H3S-2014-PDF#9(1).pdf

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Department
Biological Sciences
Course Code
BIOB12H3
Professor
Aarti Ashok

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BIOB12H3S:  Cell  &  Molecular  Biology  Laboratory       Fluorescence  Microscopy  module  (PDF  #9)     Fluorescence  Microscopy:   Fluorescence  microscopy  is  a  very  powerful  technique,  which  is  most  often   associated  with  immunofluorescence  experiments.  In  this  technique,  small   fluorescent  molecules,  known  as  fluorophores  or  fluorescent  dyes  conjugated  to   antibodies  that  recognize  specific  cellular  proteins,  are  employed.    The  dyes  absorb   light  at  one  wavelength,  and  emit  light  at  a  longer  wavelength.    Rhodamine  (emitting   red  light)  and  fluorescein  (emitting  green  light)  are  examples,  and  they  may  be   coupled  to  antibodies  or  other  molecules,  which  interact  specifically  with  cellular   components.    In  this  way,  the  subcellular  location  and  distribution  of  specific   proteins  or  other  cellular  components  can  be  observed.    This  type  of  microscopy   differs  from  regular  light  microscopy  in  that  it  provides  us  with  a  means  of   specifically  visualizing  our  protein  of  interest  within  cells  at  a  high  resolution   against  a  dark  background  provided  by  all  other  non-­‐fluorescent  components.       The  fluorescence  microscope  (also  called  epifluorescent  microscope)  is  diagrammed   below.    Light  is  passed  through  an  excitation  filter  (1),  which  permits  only  the   desired  wavelength  (excitation  wavelength)  of  light  to  pass  to  the  specimen.    A   beam-­‐splitting  mirror  (2)  will  then  focus  this  excitation  light  on  to  the  specimen   containing  the  fluorescent  dyes.  The  fluorescent  dyes  will  become  excited  and  emit   light  at  the  emission  wavelength,  which,  will  be  allowed  to  get  past  the  beam   splitting  mirror  towards  the  eyepiece  or  objective  lens.  A  second  filter  (3)  inhibits   the  passage  of  light  at  any  wavelength  other  than  the  specific  longer  wavelength   fluorescent  light  emitted  by  the  specimen.  Note  that  by  selecting  specific  filters  for   each  excitation  wavelength  of  light,  one  can  separate  different  wavelengths  of   fluorescent  light  being  emitted  by  the  specimen  (red  and  green  for  example).         Source:  Essential  Cell  Biology,     Alberts  et.  al.,  3  edition,  Pg  8                           1 Limitations:     The  fluorescent  signal  is  not  permanent.  Indeed,  on  occasion,  the  strength  of  the   signal  can  be  seen  to  diminish  as  we  view  the  sample  under  the  microscope.  This   type  of  fading  is  referred  to  as  photobleaching  and  can  be  a  serious  limitation  of   this  type  of  microscopy.  Such  photobleaching  can  be  the  result  of  photo-­‐induced   oxidation  that  damages  the  fluorescent  molecules  over  time  (chemical  damage  or   covalent  modification).  Several  measures  are  available  to  combat  this  effect.  One   could  use  a  lower  strength  (power)  of  excitation  light,  but  this  typically  results  in   decreased  signal  sensitivity  as  well  (i.e.  the  fluorescent  signal  is  very  weak).  More   photostable  dyes  have  been  developed  (compared  to  the  original  fluorescein  and   rhodamine  dyes),  that  resists  the  damaging  effects  more  robustly.  Reagents  such  as   anti-­‐fade  chemicals  can  also  be  applied  over  the  specimen  during  the  final  stages  of   sample  preparation.  Also  note  that  cell  or  tissue  specimens  can  be  rather  3-­‐ dimensional,  with  cells  present  in  multiple  focal  planes  (or  depths)  in  the  specimen.   An  average  of  the  emitted  light  from  these  multiple  focal  planes  reaches  the   objective  in  these  microscopes  and  hence  can  result  in  blurred  images.  This   limitation  is  overcome  through  use  of  the  confocal  microscope,  which  we  will   discuss  later.     Using  Fluorescence  Microscopy  to  examine  biological  specimens:   One  of  the  simplest  ways  to  examine  specimens  in  a  fluorescence  microscope  is  to   use  the  property  of  autofluorescence  of  certain  types  of  specimens.  In  this  case,   samples  such  as  pollen  grains  or  even  some  human  and  plant  tissues,  for  instance,   have  an  inherent  ability  to  emit  fluorescence  (or  react  to)  in  a  wide  range  of   wavelengths  that  can  be  captured  in  images  using  an  epifluorescent  microscope,   without  having  any  fluorescent  dyes  attached  to  them.  This  is  however  not  a  reliable   means  of  identifying  specific  features  of  these  specimens  but  can  be  used  to   understand  the  general  structure  or  layout  of  that  tissue  or  sample.  The  image   below  shows  autofluorescence  of  a  plant  stem  section  under  multiple  wavelengths   of  fluorescence  illumination.  The  red  autofluorescence  is  emitted  by  chloroplasts,   while  lignin  (a  component  of  cell  walls)  is  revealed  by  the  blue  autofluorescent   signal.  While  these  signals  do  not  allow  us  to  identify  specific  proteins,  we  are  able   to  understand  the  overall  structural  arrangement  of  this  tissue.                 Source:  http://www.noble.org/AppFiles/ImageGallery/CellImageGallery/Images/Image11.jpg   2 Specific  detection  of  macromolecules  (commonly  protein,  but  DNA  and  RNA  can  also   be  visualized)  typically  requires  the  use  of  fluorophores  such  as  dyes  (Fluorescein,   rhodamine,  commercially  available  Alexa-­‐Fluor  based  dyes  etc.)  or  fluorescent   proteins  (GFP,  RFP,  YFP  etc.).         Direct  visualization  of  fluorescent  proteins:   Fusion  proteins  can  be  made  by  fusing  the  gene  that  encodes  for  your  protein  of   interest  with  the  gene  that  encodes  a  fluorescent  protein,  such  as  GFP  (green),  RFP   (red),  YFP  (yellow)  etc.  These  recombinant  DNAs  or  constructs  can  then  be   transfected  into  cells  in  order  to  express  the  fluorescently  tagged-­‐protein  of  interest   (see  diagram  below).  Using  an  epifluorescent  microscope,  one  can  then  visualize  the   location  of  the  tagged  protein.           Adapted  from  Source:   http://bsp.med.harvard.edu/sites/bsp.med.harvard.edu/files/Image/student_resources/dna_to_protein.gif     Fluorescent  Antibody-­based  visualization  of  proteins:   This  approach  is  referred  to  as  immunofluorescence  and  involves  the  use  of   fluorescent  antibodies  to  detect  specific  proteins  within  cells  and  is  the  most   commonly  used  technique  in  conjunction  with  the  epifluorescent  microscope.  The   antibodies  are  usually  tagged  to  a  fluorescent  dye  such  as  fluorescein  or  any  of  a   number  of  commercially  available  dyes  that  can  be  excited  at  specific  wavelengths.   When  an  antibody  that  is  conjugated  to  such  a  fluorophore  is  added  to  cells,  it  will   bind  to  its  target  protein  (your  protein  of  interest).  Excitation  of  the  fluorophore   conjugated  to  the  antibody  would  therefore  reveal  the  location  of  the  protein  of   interest.  This  antibody  is  referred  to  as  a  primary  antibody  as  it  directly  interacts   with  the  protein  of  interest  and  this  method  is  known  as  direct   immunofluorescence.  The  main  advantage  of  this  approach  is  that  there  are  fewer   steps  in  the  staining  procedure  and  it  is  therefore  a  faster  technique  than  the   indirect  method  that  we  will  discuss  below.  The  tagging  of  the  primary  antibodies   with  the  dyes  (as  described  above)  uses  a  simple  chemical  conjugation  reaction  that   links  the  dye  to  a  specific  amino  acid  side  chain  on  the  antibody.  While  this  is  a   relatively  efficient  technique,  it  does  require  that  a  significant  amount  of  the   primary  antibody  be  available  for  the  in  vitro  chemical  conjugation  reaction.    In   3 addition,  it  does  require  significant  effort  on  the  part  of  the  research  scientists   carrying  out  the  conjugation  reactions.     If  the  primary  antibody  is  not  abundantly  available,  or  a  fluorescently  tagged   primary  antibody  is  not  readily  available  commercially,  then  a  different  approach,   known  as  indirect  immunofluorescence  is  often  utilized.  In  this  method,  a   secondary  antibody  that  binds  to  the  primary  antibody  is  purchased  from   commercial  sources  (in  vast  quantities)  and  used  in  the  chemical  conjugation   reactions  to  produce  fluorescently  labeled  secondary  antibodies.  In  the  actual   staining  procedure  then,  the  unlabeled  primary  antibody  is  added  to  cells,  which   then  binds  to  the  protein  of  interest.  This  is  followed  by  the  addition  of  the   fluorescently  conjugated  secondary  antibody,  which  will  bind  to  the  primary   antibody  (already  bound  to  the  protein  of  interest)  in  the  cells.    Excitation  of  the  dye   conjugated  to  the  secondary  antibody  then  allows  us  to  indirectly  locate  the  protein   of  interest  within  cells  (by  detecting  the  location  of  the  secondary  antibody,  which  is   bound  to  the  primary,  which  is  in  turn  bound  to  the  protein  of  interest).  As  multiple   secondary  antibodies  are  capable  of  binding  to  a  single  primary  antibody,  this  type   of  immunofluorescence  can  also  result  in  an  enhanced  (greater)  fluorescent  signal   compared  to  the  direct  immunofluorescence  method  (see  diagram  below).               Adapted  from  Source:  http://www.dako.com/08002_03aug09_ihc_guidebook_5th_edition_chapter_10.pdf     Sample  preparation  for  Fluorescence  Microscopy:     While  tissues  and  other  sectioned  biological  samples  can  be  used  in  direct  or   indirect  immunostaining  experiments,  we  will  only  discuss  the  use  of  cultured  cells   for  these  experiments,  as  these  are  the  most  commonly  used  specimens  for   fluorescence  microscopy  and  it  pertains  directly  to  what  you  will  be  doing  in  Lab   11B.  Primary  cells  or  cell  lines  are  typically  maintained  in  flasks  or  dishes  with   appropriate  nutrient  medium  under  5%  CO  concentration  and 2 at  37°C  (we  will   discuss  the  details  of  cell  culture  in  lecture).    For  immunofluorescence  experiments,   cells  are  cultured  on  coverslips,  which  are  small  pieces  of  glass  that  provide  a   surface  for  cells  to  adhere  to  in  culture.           4   The  pictures  below  show  a  6-­‐well  cell  culture  plate  with  each  separate  compartment   being  termed  a  “well”.  The  coverslips,  shown  on  the  right,  are  the  pieces  of  glass  that   will  be  placed  into  the  wells  of  the  6-­‐well  plate  and  onto  which  the  cells  will  grow.           Source  (left):  http://upload.wikimedia.org/wikipedia/commons/7/7b/6_well_cells_culture_plate-­‐03.jpg    ce  (right):  http://thumbs4.ebaystatic.com/d/l225/m/m1GnIFRCFpnkQNVDhJ1F0Ww.jpg     Growing  cells  on  coverslips  allows  us  to  be  able  to  later  move  them  out  of  the  6-­‐well   plate  and  place  them  on  a  microscope  slide  for  visualization  under  the  fluorescent   microscope.  For  Lab  11B,  a  human  cervical  cancer  cell  line  known  as  HeLa  will  be   grown  on  these  coverslips  placed  in  6-­‐well  plates.  Here  is  a  general  outline  of  the   steps  involved  in  preparing  such  cells  for  immunostaining  and  an  overview  of  the   immunostaining  process  (which  you  will  be  performing  in  Lab  11B).       As  a  first  step,  the  cell  culture  medium  surrounding  the  cells  will  be  removed  and   the  cells  will  be  washed  with  1X  PBS  (phosphate  buffered  saline).  This  removes  any   media  or  serum  components  that  would  otherwise  interfere  with  the  experimental   procedure.  These  cells  then  need  to  be  preserved  for  our  staining  purposes  and  the   process  is  known  as  fixation.  As  the  name  implies,  during  this  process  cellular   structures  and  organelles  become  “fixed”  in  time  and  place,  such  that  we  do  not  lose   our  proteins  or  structures  of  interest  while  going  through  the  immunostaining   procedure.    A  common  method  that  is  used  to  do  this  is  to  use  aldehyde  fixatives   such  as  glutaraldehyde  or  paraformaldehyde.  These  will  crosslink  proteins  within   cells  and  preserve  subcellular  structures  close  to  their  native  conformation.   Alternatively,  organic  solvents  such  as  methanol  can  be  used  to  fix  cells.  These   solvents  act  by  precipitating  proteins  and  the  key  difference  is  that  they  also   denature  proteins,  making  it  difficult  to  detect  any  subcellular  structures  using   antibodies  that  recognize  them  only  in  a  native  configuration.  HeLa  cells  that  you   will  use  in  Lab  11B  will  be  fixed  in  a  solution  of  4%  paraformaldehyde  for  about  20   minutes  at  room  temperature.  Residual  fixative  will  be  removed  by  washing  with  1X   PBS.  This  method  is  used  in  order  to  preserve  the  cytoskeletal  network  (that  you   will  be  visualizing  by  staining)  in  a  native  configuration.       The  next  step  is  to  permeabilize  cells  such  that  the  antibodies  that  we  will  add  to  the   specimens  for  the  fluorescent  staining  procedure  can  actually  enter  into  the  cell   5 (and  subcellular  structures).  While  the  fixation  step  does  permeabilize  the  plasma   membrane  to  some  degree,  this  step  follows  on  from  that  to  provide  more  extensive   access  to  the  intracellular  structures  and  compartments.  Cells  are  permeabilized   through  incubation  in  low  concentrations  of  detergent  solutions  such  as  Triton  X-­‐ 100,  NP-­‐40  or  saponin.  The  choice  of  detergent  and  the  concentration  used  will  vary   depending  on  the  identity  of  the  protein  or  cellular  structure  that  you  are  trying  to   stain.  For  the  HeLa  cells  you  will  use  in  Lab  11B,  we  will  have  permeabilized  them   by  incubation  in  a  solution  of  0.1%  Triton  X-­‐100  for  20  minutes.       Prior  to  beginning  the  immunostaining  procedure,  a  blocking  step  is  often   performed.  In  this  step,  fixed  and  permeabilized  cells  are  incubated  with  PBS   containing  either  serum,  BSA  or  milk  which  are  all  sources  of  protein  to  block  any   non-­‐specific  antibody  binding  sites  in  your  cell  specimens.    Non-­‐specific  interactions   (such  as  ionic  interactions)  of  antibodies  with  proteins  other  than  their  specific   antigen  are  common  and  can  contribute  to  what  is  termed  as  “background”  signal  in   immunofluorescence  experiment.  The  background  in  such  cases  would  display  a   diffusely  fluorescent  signal  rather  than  the  clear  dark  background  that  is   characteristic  of  a  good  immunostaining  experiment.  For  the  HeLa  cells  you  will  use   in  Lab  11B,  we  will  have  placed  them  in  a  blocking  solution  containing  5%  fetal   bovine  serum  (FBS)  in  1X  PBS  for  an  hour  at  room  temperature  to  improve  the   signal  to  noise  quality  of  your  immunofluorescence  experiment.       Finally,  it  is  now  time  to  actually  “stain”  your  cells  with  the  antibody  solutions.  In   most  cases,  primary  antibodies  are  diluted  (range  of  1:100  to  1:2000  is  typical)  in  a   1X  PBS  solution  that  contains  about  1%  FBS.  They  are  then  carefully  added  to  the   well  containing  the  coverslips  and  incubated  for  an  hour  at  room  temperature.   Excess  primary  antibody  that  is  unbound  is  then  removed  by  extensive  but  VERY   GENTLE  washing  of  the  cells  with  1X  PBS.  In  a  direct  immunostaining  experiment   (where  the  primary  antibody  is  directly  conjugated  to  a  fluorophore),  this  would   then  be  the  end  of  the  staining  steps  of  the  procedure.  In  an  indirect   immunostaining  experiment,  however,  the  next  step  would  be  the  incubation  of  the   cells  with  a  diluted  secondary  antibody  solution  (1:500  to  1:10,000  range)  in  1X  PBS   containing  1%  FBS  for  an  hour  at  room  temperature.  Note  that  this  step  can  amplify   the  fluorescent  signal  produced  in  your  experiment  (as  described  on  page  4  of  this   PDF),  as  multiple  fluorescently  conjugated  secondary  antibodies  can  bind  to  a  single   unlabeled  primary  antibody  that  is  bound  to  the  protein  of  interest.  Excess   secondary  antibody  is  again  removed  by  extensive  and  gentle  washing  in  1X  PBS.       This  is  the  point  at  which,  the  cell  staining  procedure  is  now  complete.  In  order  to   actually  view  your  stained  sample  under  the  fluorescent  microscope,  however,  we   must  move  the  coverslips  from  the  6-­‐well  plate  on  to  a  microscope  slide.  This  is   done  by  placing  a  small  amount  of  mounting  medium  on  a  microscope  slide  and   then  placing  the  coverslip  cell  side  down  on  to  the  medium  such  that  there  is  an   even  amount  of  the  medium  trapped  between  the  coverslip  and  the  microscope   slide.  The  mounting  medium  can  contain  DNA  dyes  (such  as  DAPI,  which  can  be   visualized  as  a  blue  stain  as  its  fluorescence  emission  maximum  occurs  at  around   6 450nm),  which  allows  us  to  easily  discern  the  morphology  of  subcellular  structures   in  relation  to  the  nucleus.  The  mounting  medium  that  you  will  use  in  Lab  11B   contains  this  DAPI  dye  and  in  addition,  contains  scavenger  molecules  that  help  soak   up  any  free  radicals  in  your  sample,  thereby  reducing  photobleaching  (see  page  2  of   this  PDF  for  a  description  of  photobleaching).    Once  the  coverslip  has  dried  on  to  the   surface  of  the  microscope  slide,  the  specimen  is  said  to  be  “cured”  or  set  in  place  and   can  now  be  examined  in  a  fluorescent  microscope.  Note  that  you  will  need  to  have  a   cured  slide  in  order  to  be  able  to  place  it  coverslip  side  DOWN  on  to  the  stage  of  a   fluorescent  microscope.       This  has  now  taken  you  through  the  entire  process  of  specimen  preparation  for   fluorescent  microscopy  of  immunostained  cells.  In  the  interest  of  time,  we  will   provide  you  with  HeLa  cells  that  have  been  fixed,  permeabilized  and  blocked  in  Lab   11B.  You  will  then  be  able  to  continue  with  the  remaining  steps  of  immunostaining   and  mounting  prior  to  Lab  12A,  in  which  you  will  get  to  image  the  slides  that  you   prepared  in  Lab  11B.                       7 Lab  11A:  Imaging  the  cell  cycle  using  fluorescence  microscopy     You  will  recall  from  BIOB11H  lectures,  that  the  cell  cycle  is  defined  as  the  events   that  occur  between  one  cell  division  and  the  next.  The  phases  of  the  cell  cycle  can  be   broadly  divided  into  Interphase  (G1,  S  &  G2),  when  the  cell  is  preparing  for  division   and  M-­‐phase  (Mitosis  (Prophase  to  Telophase)  and  cytokinesis)  when  cells  actually   divide.    The  diagram  below  illustrates  the  progression  through  these  different   phases.           th       Karp  text,  6  edition,  Fig.  14.1     Movement  from  one  stage  to  the  next,  such  as  from  G1  into  S  phase,  is  controlled  by   various  cellular  factors,  including  proteins  known  as  cyclin  dependent  kinases   (CDKs)  whose  activity  level  controls  progression  through  the  cell  cycle  by   stimulating  (or  inhibiting)  the  activity  of  other  proteins  that  aid  in  processes  such  as   DNA  replication.  CDKs,  when  active,  therefore  act  on  substrates  by  phosphorylating   them.  These  cyclin  dependent  kinases  are  interesting  proteins  in  that  their  activity   relies  on  the  concentration  of  one  of  its  subunits,  the  cyclins,  at  specific  phases  of  the   cell  cycle.  Cyclin  levels  rise  and  fall  at  different  phases  such  that  those  that  control   transition  into  S-­‐phase,  have  peak  expression  levels  in  G1  and  S  phase  while  mitotic   cyclins,  which  control  entry  into  M-­‐phase,  are  present  at  high  levels  primarily   during  the  G2-­‐M  transition.  The  figure  below  shows  the  cyclic  nature  of  cyclin   expression.       8   Karp  text,  6  edition:  Fig  1   This  means  that  the  cell  controls  progression  through  the  cell  cycle  by  inducing  the   expression  of  specific  cyclins  and  then  rapidly  degrading  those  cyclins  as  necessary   for  cell  cycle  progression.    Note  that  the  cyclins  themselves  do  not  have  enzymatic   activity,  but  are  required  in  order  to  assemble  CDK  complexes,  which  are  then  active   enzymes.  The  cyclic  nature  of  expression  of  these  cyclin  proteins  is  one  example  of   several  proteins  whose  expression  and  subsequent  rapid  degradation  is  required   for  progression  of  a  cell  through  the  different  phases  of  the  cell  cycle.  Such  proteins   may  be  present  only  during  a  subset  of  phases  of  the  cell  cycle  and  observing  their   expression  could  actually  tell  us  which  phase  of  the  cycle  a  cell  may  be  in  at  any   given  time.       Recently,  researchers  have  created  GFP  or  RFP-­‐tagged  version  of  specific  cyclic   proteins,  such  that  their  expression  or  degradation  can  be  followed  simply  by   examining  their  fluorescent  signals.  For  instance,  if  a  protein  that  is  expressed   during  G1  and  the  G1-­‐S  transition,  but  is  rapidly  degraded  in  S  phase  could  be   tagged  with  RFP,  a  red  signal  would  be  observed  when  these  cells  are  in  G1  and  G1-­‐S   but  not  during  S,  G2,  M  phase  or  M-­‐G1  transition.  Similarly,  a  protein  that  is   expressed  during  the  G1-­‐S  transition,  S  phase,  G2  and  M,  but  is  rapidly  degraded  at   the  end  of  M  phase  (and  is  therefore  not  present  in  the  M-­‐G1  transition  or  in  G1   phase)  could  be  tagged  with  a  GFP,  and  a  green  signal  would  be  observed  beginning   at  G1-­‐S  transition  into  S,  G2  and  M  phases  but  would  not  be  observed  at  the  M-­‐G1   transition  and  during  G1  phase.  The  diagram  below  shows  a  pictorial  representation   of  the  fluorescent  signals  that  would  be  observed  when  these  2  different   fluorescently  tagged  proteins  are  expressed  in  cells  together.  Note  that  during  the   G1-­‐S  transition,  cells  would  appear  yellow  (or  orange)  due  to  the  overlap  of  the  red   and  green  signals  produced  by  the  expression  of  both  proteins.  At  the  end  of  M   phase  and  re-­‐entry  of  cells  into  prophase  (G1),  neither  fluorescent  protein  is   expressed  and  the  cells  would  have  a  very  brief  colourless  phase  at  this  point.       9         Source:  http://www.amalgaam.co.jp/products/advanced/img/fucci/fucci_image.gif     Thus  far,  we  have  referred  to  these  2  proteins,  whose  expression  levels  change  over   the  course  of  the  cell  cycle,  hypothetically.  Let  us  now  refer  to  these  proteins   specifically  as  Cdt1-­‐RFP  and  Geminin-­‐GFP.    Miyawaki  and  colleagues created  HeLa   cells  (human  cervical  cell  line  commonly  used  in  biological  research)  that  stably   express  Cdt1-­‐RFP  and  Geminin-­‐GFP,  such  that  cell  cycle  progression  of  these  cells   can  be  monitored  by  fluorescence  microscopy  (Sakaue-­‐Sawano  et  al.  (2008)  Cell   132:487–498).  These  cells  are  called  HeLa-­‐FUCCI  cells,  for  Fluorescent   Ubiquitination  Cell  Cycle  Indicator  cells.  We  will  be  using  these  very  unique   fluorescent  cells  as  a  means  to  understand  the  cell  cycle  using  the  fluorescent   microscopes  in  lab  today.         A  summary  of  the  expression  pattern  and  hence  the  colour  indicator  of  the   different  phases  of  the  cell  cycle  of  HeLa-­‐FUCCI  cells  are  shown  in  the  diagram   below.  Fluorescent  images  of  HeLa-­‐FUCCI  cells  as  they  go  through  the  cell  cycle  is   also  shown  below,  to  give  you  a  primer  on  what  your  own  cell  images  may  look  like.     10 Geminin- GFP Cdt1-RFP         Adapted  from  Source:  https://tools.lifetechnologies.com/content/sfs/gallery/thumb/s007026.jpg           RFP-Cdt1 GFP- Geminin Merge             Adapted  from  Source:  Sakaue-­‐Sawano  et  al.  (2008)  Cell  132:487–498 11 Procedure  for  Lab  11A:     1.  You  will  first  be  instructed  in  the  proper  and  CAREFUL  use  of  the  rather  delicate   and  very  expensive,  research-­‐grade  fluorescent  microscopes.  The  TAs  will  give  you   very  specific  instructions  for  the  operation  of  the  microscope  hardware  and   associated  analysis  software,  which  will  be  used  to  acquire  and  analyze  images  of   cells.  NEVER  use  the  microscope  unsupervised!       2.  Each  bench  of  students  will  be  provided  with  3  coverslips  (cell  samples)  to  image.   As  we  will  have  2  fluorescent  microscopes  per  lab,  2  or  3  groups  will  need  to  share   time  on  a  microscope.  The  TA  will  now  assign  your  bench  to  a  microscope.     3.  The  first  sample  to  be  imaged  will  be  labeled  “Untreated”  and  these  are  HeLa   FUCCI  cells  that  were  grown  on  coverslips,  fixed  and  mounted  on  microscope  slides   for  visualization.       (A)  With  the  help  of  the  TA  you  will  first  examine  the  cells  under  fluorescence  using   the  objective  on  the  fluorescence  microscope.  Ensure  that  each  member  of  your   bench  has  a  turn  looking  at  the  cells,  but  be  aware  that  extensive  fluorescence   illumination  of  your  sample  will  lead  to  the  signal  fading  (photobleaching,  as   described  under  the  Limitations  section  on  page  2).  Note  that  when  you  examine  the   cells  under  UV  light,  be  very  careful  to  NOT  stare  at  the  light  for  any  extended  period   of  time.  This  can  be  VERY  harmful  to  your  vision!   (B)  Next,  you  will  use  the  camera  attached  to  the  microscope  to  capture  images  of  2   different  fields  of  cells.     (C)  You  will  then  be  ready  to  analyze  each  image.  As  different  cells  may  be  at   different  phases  of  the  cell  cycle,  you  will  use  the  colour  of  the  cells  as  an  indicator   to  count  the  number  of  cells  in  (a)  G1,  (b)  G1-­‐S  transition,    (c)  S  phase,  
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