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Imaging Experimental Design

Imaging experimental design uses high-powered microscopy and mass spectrometry techniques to capture fine detail and real-time molecular dynamics including:

  • Structural illumination microscopy (SIM)
  • Fluorescence in situ hybridization (FISH)
  • Mass spectrometry imaging (MALDI)
  • Live cell holographic imaging on NanoLive and the lattice light-sheet microscope.

Access the workflow steps for each specific imaging experimental design process.

SIM and Airyscan provide a higher spatial resolution than that of confocal microscopy. They have similar sample requirements as those of confocal microscopy. They are on the same platform as the confocal microscope instrument, and switching between them is easy. 

Image from Structure Illumination Microscopy at cell membrane level
Image produced using structure illumination microscopy highlighting the cell membrane. (Image provided by Dehong Hu | Environmental Molecular Sciences Laboratory)

1. Sample Preparation 

  • Collect live cells or tissues with intrinsic fluorescence (fluorescent proteins, pigments) or stained samples (dyes, immunostains, fluorescence in situ hybridization (FISH), activity-based probes, etc.) 
  • Mount sample using common microscopy tools (microscope slides, coverslips, glass bottom dishes) 

2. Sample Imaging 

  • Image at 10X to 100X magnification 
  • Use SIM or Airyscan mode with high magnification objectives for samples requiring a higher resolution 

3. Image Processing 

  • Use the tools in ZEN software 
  • Adjust and compile images in free open-source software: ImageJ 
  • Optionally, use third-party commercial software: Volocity 
Contact

Dehong HuDehong.Hu@pnnl.gov

The NanoLive holotomography microscope for live cell imaging is an automated platform for the observation of the same cells on the order of minutes to days, with three-channel fluorescence and multiple loading formats. 

illustration of the light path used for tomography and fluorescence data collection on the CX-A Nano Live instrument
NanoLive was used to create a timecourse of Neurospora crassa growth. Distribution of a membrane-linked enzyme which methylates ceramide lipids, scumbo, is tracked by a transcriptional fusion with green fluorescent protein (upper panel).  (Microscopy image provided by Erin Bredeweg)

1. Sample preparation 

  • Precultivate strains and treatments on site; limit cell density and z-plane growth: 
    • Bulk collection from fluorescence activated cell sorting
    • Axenic or mixed cultures 
  • Load samples in chosen format: 
    • 60-well plate (90 µL) 
    • 4-well dish format (1.5 mL) 
  • Apply dyes or stains 
  • Allow time for temperature to reach equilibrium before imaging 
  • Treat samples to obtain a flat liquid surface (60-well format) 

2. Initial Cell Population Analysis 

  • Analyze population by cell size/complexity 
  • Sort subset on gate criteria 
  • Verify expectation/accuracy of sorting by confocal microscopy 

3. Sorting 

  • Isolate single cells into 96/384-well formats 
  • Collect bulk subpopulation(s) 
  • Quantify abundance or proportions as an endpoint 
Contact

Erin Bredewegerin.bredeweg@pnnl.gov

Lattice light-sheet live microscopy increases the speed of in situ three-dimensional fluorescence microscopy with reduced damage to live cells caused by phototoxicity and photobleaching. 

Lattice-Light Sheet fluorescent image of Nitzschia inconspicua. The red is chlorophyll. The green is dye-stained lipid droplet.
Lattice light-sheet microscopy an optical lattice to create an ultra-thin light sheet that enables the imaging of whole organisms, like plants, down to single molecules using extremely low photo-dosage and phototoxicity. In this fluorescence image, lattice light-sheet was used to imageNitzschia inconspicua. The red color is chlorophyll and the green is dye-stained lipid droplets.

1. Sample Preparation 

  • Grow cells on  on 5-mm glass coverslips or mount on after culturing (either at the user’s facility or in EMSL) 
  • Larger live samples (e.g., tissues, roots, leaves) can be directly mounted onto the sample holder 
  • Cells are stained by fluorescent dye, fluorescent antibody or FISH. Cells with intrinsic fluorescence can be imaged directly.
  • Image samples by confocal microscopy to confirm samples are in good condition 

2. Sample Imaging 

  • Prior to imaging, immerse the samples in an aqueous buffer or growth medium 
  • Perform 3D imaging using four lasers (405, 488, 561, and 640 nm) as fluorescence excitation sources  
  • For time-lapse imaging, set parameters for a defined period ranging from minutes to hours 
  • 3D STORM microscopy can be used for super-resolution imaging 

3. Image Processing 

  • Visualize 3D images with Slidebook software 
  • Process 3D STORM data with an analysis program in MATLAB 
Contact

Dehong HuDehong.Hu@pnnl.gov

Fluorescent In Situ Hybridization (FISH) is a probe-based technique to visualize DNA or RNA at the single-cell level to identify functional and/or phylogenetic features and to quantify mRNA expression levels with a high spatial resolution. 

1. Sample Preparation 

  • Collect samples (cells or tissues) and store at −20 to −80°C 
  • Cryosection thick tissues and large samples 
  • Apply a fixative or solvents to preserve RNA integrity 
    • Use PFA/ethanol depending on the sample type and freezing timeline 
    • Perform PFA fixation after sectioning 

2. Design of FISH Probes 

  • Select the appropriate FISH method based on RNA abundance and the fluorescence background level of the samples: 
    • catalyzed reporter deposition (CARD-FISH) 
    • hybridization chain reaction (HCR-FISH) 
    • single molecule (smFISH) and fluctuation localization imaging-based (fliFISH) for mRNA counting 
  • Identify FISH targets (i.e., mRNA, rRNA) and specific nucleotide regions for hybridization 
  • Design oligonucleotide probes 

3. Hybridization 

  • Permeabilize cell membrane/cell wall 
  • Hybridize probe and wash 
  • Amplify signal if required (CARD-FISH and HCR-FISH) 

4. Imaging* 

  • Confocal microscopy for most samples 
  • SIM/Airyscan for a higher spatial resolution than that of confocal microscopy 
  • STORM for single transcript counting with smFISH and fliFISH 

* See specific workflows for image analysis details 

CONTACT

Dehong HuDehong.Hu@pnnl.gov

MALDI-MSI can be used to visualize the distribution of biomolecules (proteins, peptides, lipids, metabolites) on substrate and tissue surfaces. 

1. Sample Preparation & Treatment 

  • Prepare flash-frozen tissue or cell samples without chemical fixation by using polymer embedding (hydroxypropyl methylecellulose and polyvinylpyrrolidone, carboxymethyl cellulose, etc.) or cryosectioning 
  • Prepare formalin-fixed paraffin-embedded samples by microtome sectioning 
  • Remove unwanted molecules and contaminants (e.g., salts, lipids, wax) by washing in the appropriate solvent
  • Perform on-tissue chemical derivatization if needed
  • Apply MALDI matrix (automatized spraying, sublimation) 

2. Mass Spectrometry Image Acquisition 

  • MALDI-12T Fourier-transform ion cyclotron resonance (FTICR), MALDI-trapped ion mobility time-of-flight mass spectrometry (timsTOF), MALDI-ultra-high mass range (UHMR)
  • Scan slides, defining regions of interest and selection of step (pixel) size (10 µm–200 µm)
  • Run Imaging (~1pixel per s for MALDI-FTICR and MALDI-UHMR, and ~10 pixels per s for timTOF)

3. Image Analysis 

  • Upload raw data to SCILS, Bruker lab software for mass spectrometry analysis 
  • Create and upload an imzML file to the METASPACE software platform for molecular annotation and visualization 
  • Perform a statistical analysis in SCILS (receiver operating characteristic curve graphs for probability, principal component analysis, colocalization analysis, registration of microscopy images, discriminant peaks, etc.) 
Contact

Dušan Veličkovićdusan.velickovic@pnnl.gov