2011

Table of Contents

1. What is Turku BioImaging? 12. A brief history of Turku BioImaging 33. Turku BioImaging facilities 54. Light microscopy 105. Confocal microscopy 126. High content analysis 167. STED microscopy 198. FRET microscopy 229. FCS and RICS 2410. Two-photon microscopy 2611. Spinning disk confocal microscopy 2912. TIRF microscopy 3113. ScanR automated fluorescence microscopy 3314. Stereo microscopy with microinjection 3615. Time-resolved fluorescence microscopy 3816. Live cell imaging 4117. Image analysis 4418. Flow cytometry 4719. Cell sorting 5020. Flow cytometry data analysis 5221. Atomic force microscopy 5422. Electron microscopy 5723. PET imaging 6024. Small animal ultrasound imaging 6225. Micro-CT imaging 6426. Imaging mass spectrometry 6627. Macroscopic in vivo imaging 6828. Laser-capture microdissection 7029. Master’s Degree Programme in Biomedical Imaging 7230. Turku as an imaging stronghold 75Turku BioImaging email addresses 77

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1. What is Turku BioImaging?

Turku BioImaging is an interdisciplinary imaging organization that aims to bring together bioimaging expertise in Turku.

Turku BioImaging consortiumTurku BioImaging was initiated as a joint program of the Univer-sity of Turku and Åbo Akademi University. The organization is a broad-based science and infrastructure umbrella, and aims to unite bioimaging expertise in Turku and elsewhere in Finland. The purpose is to maximize synergism, to engage in complemen-tary activities, and to promote both regional and national devel-opment of imaging technologies and instrumentation.

Turku BioImaging embraces state-of-the-art imaging technolo-gies used by the bioscience community of Turku. The organi-zation is highly interdisciplinary, encompassing a wide range of imaging modalities from molecular to cellular, from single molecule analysis to whole animal imaging, and from analysis of events at the single cell level to high-throughput screening. The initiative also includes proteomics, systems biology, and computational modeling of cellular processes, each of which is associated with specific bioimaging modalities. Further, Turku BioImaging also offers software tools for image data processing and analysis. Turku BioImaging is closely integrated with the Biocenter Finland National Imaging Infrastructure Network (http://www.biocenter.fi/index3_biologicalimaging.html), as well as with the pan-European imaging infrastructure EuroBioimaging (www.eurobioimaging.eu).

MSc Programme in Biomedical ImagingThe various university departments and core facilities included in Turku BioImaging have a great deal of experience in provid-ing training in imaging and in facilitating access to modern equipment. In this regard, the latest development is a novel in-ternational Master of Science Degree Programme in Biomedical

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Imaging. The MSc programme, jointly administered by the Uni-versity of Turku and Åbo Akademi University (please see chapter 29), spans all imaging modalities and features lectures from local and international experts in bioscience, medicine, physics, com-puter science, proteomics, and the visual arts.

More information: www.bioimaging.fi

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2. A brief history of Turku BioImaging

The Turku BioImaging consortium was initiated in 2007 as an interdisciplinary co-operative project between the two universi-ties in Turku. The aim of the association is to promote expertise in biomedical imaging in the Turku region. The Turku BioImag-ing Kick-off Seminar was held in BioCity Turku on 11 June 2007.Since that time, Turku BioImaging has become a key feature of the Turku university campus and serves to co-ordinate the devel-opment of molecular imaging within the Turku region. The goals of Turku BioImaging include development of interdisciplinary research projects and the establishment of a continuum of imag-ing techniques, ranging from experimental concepts to clinical and pharmacological applications.

In early 2009, the Ministry of Education selected the Turku Bio-Imaging consortium as one of the 24 most significant national-level research infrastructures in Finland (Finnish research in-frastructure roadmap: http://www.minedu.fi/export/sites/default/OPM/Julkaisut/2009/liitteet/opm02.pdf?lang=fi).

Also in 2009, the key stakeholders signed an agreement on the future of Turku BioImaging, with the aim of creating a lead-ing bioimaging technology platform in Northern Europe in the coming years. The partners of the initiative are the University of Turku; Åbo Akademi University; Turku Science Park; the Hospi-tal District of Southwest Finland; the Centre for Economic De-velopment, Transport, and the Environment of Southwest Fin-land (the ELY Centre); and the Regional Council of Southwest Finland.

BioCity, the central building of Turku BioImaging.

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At present, Turku BioImaging is extensively involved in coordi-nation of the National Imaging Research Infrastructure Network of Biocenter Finland, and participates actively in the EuroBio-imaging initiative for the development of biomedical imaging infrastructure in Europe.

Cooperation agreement for the future of Turku BioImaging. A Signature Ceremony was held on September 30, 2009, at the Mauno Koivisto Centre in BioCity, Turku.

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3. Turku BioImaging facilities

Turku BioImaging brings together highly successful imaging facili-ties, and taps into various areas of imaging expertise on the Turku university campus.

Turku BioImaging is an umbrella organization encompassing all imaging resources on the Turku university campus. Within the co-operative Turku BioImaging consortium, both universi-ties in Turku have joined forces to establish an imaging network for development of bioimaging expertise in the Turku region. Turku BioImaging includes the national Turku PET Centre and the Cell Imaging Core (CIC) at the Turku Centre for Biotechnol-ogy. These are both joint facilities of the University of Turku and Åbo Akademi University. Turku BioImaging also includes the fa-cilities of the Turku Center for Disease Modeling, the University of Turku Electron Microscopy Unit, and imaging-related facili-ties of the VTT Technical Research Centre of Finland in Turku. Moreover, the multidisciplinary network of Turku BioImaging integrates closely with research programs in systems biology, proteomics, and information technology.

Turku Cell Imaging CoreThe Cell Imaging Core (CIC) is a centralized facility where us-ers receive the support needed to use both conventional and advanced imaging techniques. The unit is a combination of resources from Åbo Akademi University and the University of Turku. The facility has state-of-the-art imaging and flow cytome-try equipment, which are available to all researchers. In addition, the CIC provides hands-on training to users and offers consulta-tion on experimental design and image analysis. For experienced

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users, the CIC organizes advanced workshops on imaging and flow cytometry methods and applications. The CIC is repre-sented in the Biocenter Finland Bioimaging Infrastructure, and maintains links with national and international imaging centers, and coordinates the Nordic Imaging Network. http://cic.btk.fi/.

Turku PET CentreThe Turku PET Centre integrates high-level scientific expertise with excellent facilities and equipment located centrally on the university campus and in the university hospital. The Centre offers PET expertise for drug discovery and development and clinical work linking pathophysiology and disease. Human PET imaging facilities are available, and animal PET imaging may be performed in co-operation with the Turku Center for Disease Modeling. The Turku PET Centre has a partnership agreement with Turku Imanet Oy (GE Healthcare; Medical Diagnostics).http://www.turkupetcentre.fi/

Turku Center for Disease ModelingThe Turku Center for Disease Modeling (TCDM) is a research infrastructure at the Institute of Biomedicine, University of Turku, and has advanced expertise in disease modeling using ex-perimental animals. In co-operation with the Turku PET Centre,

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TCDM offers small animal imaging services that support both academic and industrial non-clinical research (basic, preclinical, and translational). The following techniques are available: opti-cal imaging, MRI, PET, ultrasound imaging, and X-ray imaging. In co-operation with the Institute of Microbiology and Patholo-gy, TCDM offers services including imaging of tissues, detection of cellular and subcellular structures by basic histology, and im-munohistochemistry using both light and electron microscopy. http://www.tcdm.fi/

EM Core FacilityThe Laboratory of Electron Microscopy serves as a core facility for nanolevel imaging and offers transmission electron micros-copy, scanning electron microscopy, atomic force microscopy, scanning tunneling microscopy, immunocytochemistry, and elemental analysis and localization by combination X-ray spec-trometry. Samples can be prepared from tissues, cells, bacteria,

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viruses, nanoparticles, other particles, macromolecules, and many other materials. Images are recorded on sheet film or as digital image files. The laboratory offers a full service, including assistance in the planning of experiments, sample processing, microscopic examination, recording of images, and reporting and interpretation of results. http://med.utu.fi/em/

Imaging software developmentImaging methodologies produce vast quantities of data that must be transformed into formats that scientists can easily visu-alize and analyze. Thus, bioimaging is very closely allied with information technology. The processing and analysis of bioim-aging data requires the development of imaging software and computational modeling. BioImageXD (please see chapter 17) is a free open-source software developed in collaboration by the Universities of Turku and Jyväskylä, Åbo Akademi University, and other partners. The software is available to all users of Turku BioImaging (www.bioimagexd.net). Flowing Software is a free soft-ware for Flow Cytometry data analysis (please see chapter 20). It has been developed at the Cell Imaging Core, Turku Centre for Biotechnology, and is available to all users of Turku BioImaging

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(www.flowingsoftware.com). Turku Bioimaging also administrates Asimov, a resource management software system for imaging in-struments. The system enables electronic reservation of instru-ments, invoicing and real-time statistics.

Other imaging possibilitiesIn addition to the larger facilities and centers mentioned, Turku BioImaging includes many other units offering various addition-al imaging techniques and services. These include atomic force microscopy, imaging mass spectrometry, and many specialized variations of light microscopy.

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4. Light microscopy

Light microscopy includes both basic and advanced methods whereby light is used to visualize objects. The range of applications varies from single molecules and cells to animals (e.g., worms or zebrafish). The use of fluorescence in addition to “normal” light al-lows visualization of elements that can be labeled with antibodies or fluorescent tracers. Light microscopy provides numerous critical tools for life science studies.

How does it work?Light microscopy uses ultraviolet, visible, or infrared light to de-tect objects. Because light is highly versatile, photon detection, used as an imaging modality, remains clearly separate from spe-cific imaging modalities employing electrons (EM), X-rays (CT), radioactive decay (PET), and molecular resonance in a magnetic field (MRI).

What can be studied with this technique?In all imaging, contrast is needed for visualization. Light mi-croscopy techniques include bright-field, phase contrast, and differential interference contrast (DIC) microscopy, wherein contrast is obtained either by using a phase shift, polarization, or staining with chemicals yielding visible colors. Fluorescence is not necessary. However, fluorescence is a very efficient method for production of contrast. For fluorescence imaging, Turku Bio-Imaging hosts several fluorescence and confocal microscopes (chapters 5, 6, 10, 11, 15, 16, 28, a total internal reflection micro-scope (chapter 12), and a super-resolution microscope (STED;

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chapter 7). Often, phase contrast or DIC can be combined with fluorescence imaging to gain information from both modalities. The properties of the sample material and the resolution needed define the most suitable method.

Technical detailsNormally, bright-field and phase contrast samples are visualized using a halogen lamp. When working with fluorescence, the light source can be a laser or a mercury (HBO), xenon (XBO), or met-al halide lamp. When working with lamps, a specific filter cube including three filters (excitation, dichroic mirror, and emission) is required for each fluorophore. If lasers emitting coherent light are used, excitation filters are unnecessary. The light is usually collected using color or black-and-white (b/w) CCD cameras or photomultiplier tubes (PMTs). Color CCD cameras are em-ployed primarily for imaging of stained tissue, whereas b/w cam-eras are used for fluorescence imaging because of their higher sensitivity and wider range of detection wavelengths. PMTs are the primary detector type employed in confocal microscopy.

Availability within Turku BioImagingTurku BioImaging provides a wide range of light microscopes ranging from fluorescence stereo microscope for macro imag-ing to a super-resolution confocal microscope. Most of the tech-niques available are introduced in this booklet. The responsible operators can help in determining the best method for each ap-plication, and can be contacted through the common email ad-dress below.

ContactLM@bioimaging.fi

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5. Confocal microscopy

Confocal microscopy is a light microscopy technique that can pro-duce true three-dimensional images of biological samples without harming the cells, tissue or organism. Combined with advanced software, confocal microscopy is one of the most popular and ver-satile biomedical imaging methods available.

How does it work?Confocal microscopy is a fluorescence light microscopy tech-nique whereby image detection is achieved through a small pin-hole. The image is illuminated point-by-point with a laser that excites fluorescence in the sample, and the detected light passes through a pinhole located on a confocal plane on the other side of the lens, defined by reference to the focal plane of the sample. The pinhole excludes most of the light coming from either be-low or above the focal plane. Thus, an optical section is created through the sample. After acquiring the image from one focal plane, the focus can be changed and a new image acquired. In this way a stack of optical sections covering the entire sample is obtained. This stack forms a three-dimensional image of the en-tire sample, as opposed to the two-dimensional images obtained using regular light microscopes. Because sectioning is optical, non-invasive and relatively fast, living cells can be imaged and four-dimensional images (three spatial dimensions plus time) can be successfully acquired. Specialized software can be used to create both three-dimensional images and four-dimensional movies of imaged material, as well as to perform many types of quantitative analysis.

The principle of confocal microscopy (simplified). The pinhole allows only light emanating from the focal plane (yellow) to pass through to the detector. Light coming from below (purple) or above (cyan) the focal plane is excluded.

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What can be studied with this technique?Confocal microscopy can be used for numerous purposes, ranging from localization of particular proteins inside cells, to studies of protein dynamics and interaction. Several variations on the basic imaging technique exist. For example, part of the fluorescence can be bleached, and the recovery and dynamics of the fluorescence monitored (the FRAP technique). Also, co-localization of two different fluorescent markers can be studied to reveal possible biological interactions, which can in turn be confirmed in fluorescence resonance energy transfer experi-ments (the FRET technique, chapter 8). Using recombinant gene technology, almost any protein can be made to fluoresce, and protein behavior and functionality can be studied in three or four dimensions (three spatial dimensions plus time) in either living or fixed cells.

When is this technique the right choice?Confocal microscopy can be used with virtually all fluorescent samples, and should be employed whenever three- or four-dimensional images are required. The technique is highly ver-satile and can reveal a wealth of information, but is also rather imprecise and error-prone. This means that the results should be checked with many different experiments and validated with other research methods.

What is required?Fluorescent samples prepared using suitable high-quality cover glass (or appropriate live cell containers) are required. The mi-croscopes are more complicated than basic light or fluorescence microscopes, but basic training will overcome this problem. Special software, such as BioImageXD developed by the Univer-sity of Turku (please see chapter 17), can be utilized to generate

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images and to derive quantitative results from image material. Because confocal microscopy is inherently a digital technique, a basic understanding of digital images and the processing thereof is needed.

How to get started?Getting started is easy because there are many microscopes and attendant experts available within Turku BioImaging. It is best to contact an experienced operator as a first step (see below). Courses are also run regularly.

Availability within Turku BioimagingTurku BioImaging offers several confocal microscopes, each tai-lored to slightly different applications. Responsible operators are listed for each device and can be contacted through the common email address below.

Zeiss LSM510 combined with an atomic force microscope• BioCity, floor -1, staircase A / Pasi Kankaanpää.

Leica TCS SP5 MATRIX (HCS A)• Used for high throughput and more common multi-well ap-

plications.• BioCity, 2nd floor, room 2209 / Jari Korhonen.

Leica TCS SP1• Excellent for training.• BioCity, 2nd floor, room 2077 / Jari Korhonen.

Zeiss LSM510 META• Serves as the main work horse of the Cell Imaging Core.• BioCity, 5th floor, room 5149 / Markku Saari & Jouko Sand-

holm.

Examples of confocal microscopy images created with BioImageXD software (chapter 17). On the left, internal green actin filaments & surface-located red integrins can be seen. On the right, various membrane structures differ in surface details.

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Leica TCS SP5• High resolution STED capability.• BioCity, 5th floor, staircase A / Sami Koho.

Leica TCS SP5 MP multiphoton system• BioCity, floor -1, staircase A / Tibor Veres.

For the related technique of spinning disk confocal microscopy, please see chapter 11.

Contactconfocal@bioimaging.fi

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6. High content analysis

High content analysis allows simultaneous imaging and analysis of large numbers of fixed or live microscopy samples. A typical ex-periment consists of three steps: sample preparation, automated imaging of samples, and image analysis. This method ideally pro-duces data reflecting the physiological parameters of cells in a given population. Typical assays explore the effects of siRNA or feature small molecule (drug) screens. The method is also suitable for more common multi-well plate applications. Scanning 60 wells, four fields per well with autofocusing, takes roughly one hour.

How does it work?Samples (fixed or live cells/tissues), placed on multi-well plates after different treatments, are automatically imaged by a light microscope either once, or on many occasions over time. The images can be visually analyzed or (automatically) examined by image analysis software to detect quantitative differences among samples.

Numerous areas in each well of a multi-well plate can be automatically scanned.

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What can be studied with this technique?Many properties can be analyzed, including distribution and ex-pression of a fluorescent label within cells over time, cell shape and movement, cell count and extent of apoptosis, and organelle/cytoskeleton characteristics. These parameters can be simultane-ously evaluated in numerous samples, depending on the number of wells scanned.

When is this technique the right choice?High content analysis is useful when multiple properties of cells can be observed most easily by light microscopy, and/or when it is impractical to manually process the number of samples to be imaged and analyzed. Even if the number of samples is suit-able for manual operation, this method is still worth considering whenever data from several wells of a multi-well plate is needed. Up to 60 wells can be imaged in one run, with autofocusing pro-viding in-focus images.

What is required?The requirements of high content analysis vary depending on the scale of the experiment. A bench-scale functional assay, which preferably can be performed semi-automatically, is needed. The cells analyzed with imaging software must be fluorescently stained to allow segmentation of single cells (i.e. identification of cell areas). Also, the properties to be measured must be suitable for robust quantification by automated image analysis methods. The imaging system can be a confocal or wide-field fluorescence microscope with a motorized stage and inbuilt software, allow-ing automated imaging of multiple positions and autofocusing (automatic determination of the correct focal plane). If automat-ed analysis is to be used, software suitable for analysis of imaging

Professor John Eriksson and senior researcher Pasi Kankaanpää beside a Leica TCS SP5 confocal microscope in BioCity.

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data is required. Also, depending on the scale of the experiment, a program that can handle large amounts of data and that can perform data mining may be needed. The scale of the experi-ment can also be more modest. Specimens in dishes 35 mm in diameter or on microscope slides can also be scanned.

Availability within Turku BioImagingLeica TCS SP5 MATRIX (HCS A)• BioCity, 2nd floor, room 2209.

The Matrix software allows imaging of multi-well samples auto-matically after the operator has constructed a scanning protocol. Due to technical restrictions, in a 96-well plate the outermost rows and columns cannot be used, leaving 60 wells in use. The microscope is equipped for live cell imaging (stable temperature, autofocus, and control of CO2 level). Fixed samples can also be scanned.

ContactJari Korhonen, HT@bioimaging.fi

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7. STED microscopy

Stimulated Emission Depletion (STED) microscopy is a confocal-like microscopy technique, in which the traditional diffraction resolution limit of optical microscopy can be effectively surpassed. With a STED instrument, it is possible to acquire purely optical super-resolution images, with details visible at the level of tens of nanometers.

How does it work?As first realized by Ernst Abbe in the late 19th century, the reso-lution of a standard far-field light microscope is limited by dif-fraction to about one-third of the wavelength used, which is ap-proximately 180 nm for visible light. Recent research, pioneered in the mid-1990s by Stefan Hell, has shown that the fundamen-tal diffraction barrier can be broken in fluorescence far-field microscopy, making it possible to achieve a resolution close to the scale of individual molecules. The basic principle of STED microscopy is the use of two lasers working in a synchronized manner, in a setting that is otherwise rather similar to that of scanning confocal microscopy. One of the lasers is employed for normal fluorescence excitation, whereas the other forms a red-shifted donut-shaped beam (of zero intensity in the center), which is overlaid on the diffraction-limited fluorescence spot, and then used to deplete all fluorescence at that spot, except for the immediate center. This is achieved by exploiting an optical process termed stimulated emission depletion. By scanning the sample using a series of sub-diffraction spots, a purely optical super-resolution image can be constructed.

The principle of STED.

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What can be studied with this technique?An STED instrument can be used in a manner very similar to that employed when other fluorescence far-field microscopes are used. Compared with, for example, electron microscopy, STED has the obvious benefit of being able to non-invasively image (living) specimens in three dimensions. Also, highly specific fluorescence labeling can be of great assistance, especially in the comparison of different biological samples.

When is this technique the right choice?STED is the instrument of choice when the resolution of normal confocal microscopy is insufficient.

What is required?Samples for the STED instrument are prepared in the same man-ner as for normal fluorescence microscopy. However, two con-siderations arise:• The Leica TCS STED currently has two super-resolution chan-

nels. For fluorescence labeling ATTO647N and Chromeo494

Crimson fluorescent microbeads imaged by STED and confocal microscopy.

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dyes are recommended. Antibody-conjugated versions of these labels are available.

• Because of the higher optical resolution of STED, optical mis-match is more of an issue with this technique. Therefore, an embedding medium with a refractive index as close as possi-ble to the 1.51 of oil immersion medium should be used (TDE and Mowiol are suitable). Also, thickness-corrected cover glasses should be employed, if available.

Availability within Turku BioImagingLeica TCS STED• BioCity, 5th floor, staircase A.

ContactSami Koho, STED@bioimaging.fi

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8. FRET microscopy

Förster or Fluorescence Resonance Energy Transfer (FRET) has advanced to become one of the most popular applications in mi-croscopy and fluorescence spectroscopy. It allows the visualization of molecular interactions and dynamics in living cells. This spatio-temporal mapping of molecular activity will continue to attract major attention in the post-genomic area. Developments with this methodology to meet the demands of the post-genomic era will have a significant potential in the future.

How does it work?With Förster Resonance Energy Transfer (FRET), named after Theodor Förster, a German physical chemist, nanometer-range molecular proximities between donor and acceptor fluorophores can be detected. These are typically attached to two bio-mole-cules to detect their interaction or co-clustering. If integrated into one biomolecule they can report on its conformational state. FRET can be applied in vitro, ex vivo and in vivo. Two key re-quirements for FRET are that the donor emission and the accep-tor excitation spectra overlap and that the fluorophores are only approximately 1-10 nm apart. The extent to which FRET occurs is expressed by the energy transfer efficiency, which ranges from 0 (no FRET) to 1. However, in practice various (instrument de-pendent) FRET measures or indices are used.

What can be studied with this technique?FRET is widely used in biochemical assays for high-throughput screening under the name of HTRF® or in the TaqMan probes for RNA quantification. In cell biology FRET probes have pro-vided unprecedented insight into where molecules are active and where they form complexes.

When is this technique the right choice?There are numerous applications for FRET in biology. Most prominently, scientists are interested in quantifying molecular interactions. Similarly, learning about conformational changes can be achieved with FRET. However, the distance constraints need to be kept in mind. For large complexes, alternative ap-proaches such as FCCS (Fluorescence Cross Correlation Spec-troscopy) are a better choice when assessing interactions.

What is required?FRET can be determined by measuring the change in the fluores-cence properties of the donor or the acceptor fluorophore, such as their intensity or their fluorescence lifetime. The fluorescence lifetime is the average time a fluorophore spends in its excited state. FRET can be a process that leads to de-excitation, thus

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lowering the average fluorescence lifetime. Also the local envi-ronment of a fluorophore can affect its fluorescence lifetime. To measure FRET, more or less specific instrumentation is required. While intensity changes can be readily monitored with fluores-cence plate readers, flow cytometers and microscopes, the fluo-rescence lifetime measurement requires hardware adaptations of these instruments. In cell biology this is commonly realized with Fluorescence Lifetime Imaging Microscopy (FLIM). FLIM is very sensitive and produces images that reflect the changes of the fluorophore environment, such as pH and polarity. A third way of assessing FRET is to monitor the depolarisation (anisot-ropy) of polarized light due to FRET. This method is also the only one to measure homo-FRET, which is the energy transfer between identical fluorophores.

How to get started?Setting up FRET methods and instrumentation is a time con-suming process. Furthermore, often tailoring the biological ex-periment such that FRET can be used effectively is much more demanding and expert advice should be sought whenever pos-sible (see contact information below).

Availability within Turku BioimagingFRET can be done on all of the Cell Imaging Core’s confocal microscopes and the LSRII flow cytometer (BioCity, 5th floor). Moreover, FLIM is going to be available on some instruments, like the STED microscope (BioCity, 5th floor).

ContactDaniel Abankwa & Camilo Guzmán, FRET@bioimaging.fi

FRET reveals association between PTRF (a novel coat protein of caveolae, left) and the caveolar component caveolin-1 (middle) in vesicular structures on the plasma membrane (arrowheads) and within (arrows) BHK cells. FRET was calculated per pixel as a sensitised acceptor emission index FR, ranging from 1 (no FRET) to 10 and above as displayed on the yellow-hot look-up-table in. Scale bar 2 μm.

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9. FCS and RICS

Fluorescence Correlation Spectroscopy (FCS) and Raster Image Correlation Spectroscopy (RICS) are two techniques that analyze the fluorescence fluctuations of molecules that diffuse through a femtoliter focal volume. FCS and RICS can be performed on a con-focal microscope. They provide information on parameters such as diffusion and concentration, and can also be used to identify bind-ing events and factors.

How does it work?FCS and RICS are normally implemented via a confocal micro-scope, where the sample is excited through the objective and emitted light is collected by the same objective. Subsequently the photons are counted over time (typically seconds to minutes) on sensitive detectors, such as avalanche photodiode detectors (APD or SPAD). In a more advanced setup, these fluctuation methods can also be performed on a STED-microscope. After photons are collected, an autocorrelation analysis of the fluores-cence events is performed. The autocorrelation function analyses the self-similarity of the fluorescent signal and determines the probability of whether the fluorescence signal is emitted by the same or by a different fluorescent molecule. Next the autocorre-lation function is fitted to known mathematical models that take into account all kinds of processes that may contribute to the signal fluctuation. Thus, quantitative information is obtained on such parameters as diffusion time and coefficient, hydrodynamic radius, and average concentration. The main difference between FCS and RICS is that in FCS fluorescence fluctuations are ana-

Principles of FCS and RICS. Left: single point FCS where laser focus (red) is static but recorded fluorescence fluctuates as fluorescent molecules (green dots) transit through the focal point. Right: RICS, a scanning FCS technique. Laser is not static, but is scanning a determined area. Periodicity of the scanning is exploited to obtain additional temporal information from the scanned im-age. RICS has lower temporal resolution than FCS, while being able to average fluctuation parameters such as diffusion over a larger area.

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lyzed in a fixed focal volume, whereas in RICS the illumination volume is moved to scan a larger region of interest.

What can be studied with this technique?FCS and RICS can be applied in vitro, in cells and in tissues of living organisms due to the high diversity of fluorescent chromo-phores that can nowadays be attached to different proteins and other biomolecules. FCS and RICS provide information at the single-molecule level by averaging the behavior of many mol-ecules, thus achieving very good statistics. A RICS experiment is similar to performing many single-point FCS measurements consecutively, with the difference that the temporal resolution on each point is much lower than in FCS. Because areas are scanned in RICS, the technique can generate diffusion and con-centration maps. These maps can be superimposed on the image of the scanned area.

When is this technique the right choice?FCS and RICS can are primarily used when information about the dynamics of molecular motions is needed. They are spectros-copy techniques rather than imaging techniques. They provide information about diffusion, concentration and binding events.

What is required?As a requirement for the autocorrelation analysis, nanomolar concentrations should be used. This ensures that no more than one fluorophore transits through the confocal volume at a time. For a study of binding events, it is necessary that the mass of the complex has changed at least by a factor of 8 as compared to the unbound fluorescent molecule. Therefore protein-protein inter-actions can usually not be detected with FCS. However, a related method, Fluorescence Cross Correlations Spectroscopy (FCCS), enables this. In FCCS, both putative binding partners are fluo-rescently labeled and the coincidence of their fluorescence fluc-tuations is analyzed. Availability within Turku BioimagingLeica TCS STED• BioCity, 5th floor, staircase A.

ContactDaniel Abankwa & Camilo Guzmán, FCS@bioimaging.fi

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10. Two-photon microscopy

Two-photon microscopy is a laser-scanning microscopic technique typically used for the acquisition of three-dimensional time-lapse images of living samples. As a major advantage, it allows a greater penetration depth into tissues (several hundred microns) than what is possible with conventional confocal microscopy (approx. 80 µm), with even lower phototoxicity. This technique is thus ideally suited for intravital microscopy in living, anaesthetized animals or living, explanted tissues.

How does it work?Like confocal microscopy, two-photon microscopy allows opti-cal sectioning of fluorescently labeled samples, although on the basis of a different principle. In conventional fluorescence excita-tion a single photon (emitted by a visible-light or UV continu-ous-wave laser) absorbed by a fluorophore molecule is sufficient for its fluorescence. In contrast, two-photon excitation occurs after the near-simultaneous absorption of two photons (emitted by an infrared, femtosecond-pulsed laser), each of which car-ries only half of the energy needed. Since two-photon excitation events are rare, they only occur at places with a high density of photons, i.e. in the focal point. The result is fluorescent excitation occurring exclusively in the focal plane, not above or beneath it. Thus, optical sectioning is an inherent property of the technique, and no pinholes are needed. Signal detection is achieved by us-ing close-coupled, highly sensitive non-descanned (external) PMT detectors. The utilization of high excitation wavelengths results in great tissue penetration and excitation being confined to the focal plane greatly reduces phototoxicity.

The principle of two-photon excitation (simplified).

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What can be studied with this technique?Two-photon microscopy is ideal for visualizing the activity of cells located deep within their native tissue environment. The cells of interest ideally express fluorescent proteins (e.g. EGFP, EYFP, DsRed in transgenic animals) or they are labeled in vitro with vital dyes (e.g. CFSE and SNARF-1) before they are injected into a recipient animal (this is a common way for visualizing im-mune cells, which are easy to isolate and are able to find their original tissue location after i.v. injection). Two-photon micros-copy has been widely used to study the migration and interac-tions of immune cells during the host response to infection. The technique is also suitable for studying neuronal activity in the brain, using either brain slices or intravital preparations. Intra-cellular signaling events in cells of intact tissues can also be ad-dressed, e.g. by using fluorescent ion indicators.

When is this technique the right choice?Two-photon microscopy is the right choice if one wants to visu-alize living cells deep within their native tissue environment. It is also useful if tissue fixation and labeling is not possible (e.g. due to signal loss or unwanted changes of the sample), because

3D image of an intact mouse lymph node demonstrating the possible imag-ing depth of two-photon microscopy. Red and green fluorescent lymphocytes are visible as deep as 250 µm below the capsule (blue) at the surface of the lymph node.

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unprocessed tissue pieces can be imaged. Although most fluo-rescent proteins are suitable for two-photon excitation, the tech-nique can also be used with tissues not containing fluorescently proteins, since specific fluorescence phenomena (so-called sec-ond- and third-harmonic generation) allow the visualization of certain tissue structures (collagen fibers and muscle filaments) without labeling.

Availability within Turku BioimagingLeica TCS SP5 MP• Multiphoton system with an upright DM6000 CFS micro-

scope.• BioCity, floor -1, staircase A.

ContactTibor Veres, 2-photon@bioimaging.fi

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11. Spinning disk confocal microscopy

Spinning disk confocal microscopy offers several advantages over laser scanning confocal microscopy. Spinning disk microscopes al-low acquisition of images at very high frame rates and are therefore well suited to high-speed live-cell 3D imaging.

How does it work?Spinning disk confocal microscopes use a pair of rotating disks, which have thousands of pinholes in a spiral configuration. In a Yokogawa spinning disk confocal microscope, the second disk has microlenses which increase the illumination of the sample. When light is projected onto the disk, the holes allow passage of an array of excitation light to the sample. The whole field of view is scanned during a single exposure and an image of the sample is formed very quickly. The major difference to a scanning confo-cal microscope is that, rather than scanning with a single point, multiple points are acquired simultaneously in the spinning disk confocal microscope.

What can be studied with this technique?Spinning disk confocal microscopy is best suited for study living cells with high speed 3D imaging over time. Also conventional fixed samples can be studied. Colocalization studies and 3D re-construction are the most common experiments. The spinning disk confocal microscope at VTT Turku has 488, 561 and 647nm laser lines and 5x, 20x, 40x, 63x and 100x objectives.

Yokogawa spinning disk unit optical configuration.

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When is this technique the right choice? Spinning disk confical microscopy is the right choice when high-speed 3D live-cell imaging is needed.

What is required?Living cells on a live-cell imaging dish (Mattek or Ibidi) with standard growth media. The VTT spinning disk confocal micro-scope has an environmental chamber with heating and CO2 sup-ply. Fixed cells and standard microscope slides are also suitable.

How to get started?Contact VTT Medical Biotechnology for guidance and assis-tance.

Availability within Turku BioImagingCarl Zeiss Axiovert 200M with Yokogawa CSU22 spinning disk unit• VTT Turku.

ContactAntti Arjonen, spinning@bioimaging.fi

GFP-Histone2B labeled MDA-MB-231 cells under cytokinesis (separation of sister chromatids), 4D imaging.

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12. TIRF microscopy

Total Internal Reflection Fluorescence (TIRF) microscopy is the op-timal choice for researchers interested in events that occur on the cell membrane. In TIRF microscopy, an evanescent wave caused by total reflection of the light beam is used for sample excitation. The evanescent wave itself penetrates only approximately 100 nm into the sample.

How does it work?At a specific high angle, dependent on the refractive indices of glass and water, the light from the laser source is totally reflected from the glass/water interface. A small proportion of this reflect-ed light travels parallel to the surface plane and generates a thin electromagnetic field in the aqueous phase. This electromagnetic field is termed the evanescent field and can be used to excite fluo-rophores located at or near the basal membrane of a cell.

What can be studied with this technique?Turku BioImaging hosts an Olympus CellR fluorescence micros-copy system equipped with TIRF. The instrument is suitable for high-resolution detection of events and structures in the imme-diate vicinity of the object glass-water boundary (i.e., the basal plasma membrane of the cell). The CellR system allows live cell fluorescence imaging at a time resolution of milliseconds.

When is this technique the right choice?TIRF is especially appropriate for research on phenomena at the cell surface, such as internalization and transport of vesicles, and cell adhesion. The fast camera of the CellR system also allows visualization of calcium signaling, and characterization thereof with FURA staining.

A live COS-7 cell transfected to express EGFP-tagged small GTPase Rab7. The images show Rab7-positive vesicles in close proximity to the plasma membrane (the TIRF channel) and throughout the cell body (the widefield channel).

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Availability within Turku BioImagingOlympus CellR TIRF• Medisiina, 2nd floor, room C204.

ContactJorma Määttä, TIRF@bioimaging.fi

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13. ScanR automated fluorescence microscopy

ScanR is a fully automated and highly flexible microscope-based screening platform for life science applications, developed by Olympus in collaboration with the European Molecular Biology Laboratory (EMBL).

How does it work?The ScanR microscope is a widefield fluorescence microscope that is designed for fully automated image acquisition and data analysis of biological samples and can handle many different for-mats, e.g. multi-well plates, slides or custom-built arrays. Image acquisition can be done with five different filters (DAPI, Alex-aFluor488, 555, 594 and 647) with autofocus. The microscope does not have a temperature and CO2 controlled chamber, so the setup is not suitable for live cell imaging.

What can be studied with this technique?The ScanR microscope is specifically developed for high-through-put imaging. It can automatically image many different formats, like multi-well plates and the Cell Spot Micro-Arrays developed at VTT. The Analysis software can then be used to define, for ex-ample, DAPI stained cells and subsequently count cells (in such applications as proliferation assays). Fluorescence intensity can be quantified for each cell (for instance to determine gene ex-pression or apoptosis). According to the manufacturer, the ScanR setup can be used for the following assays:• Cell counting• Gene expression• Intracellular transport• Translocation• Cell proliferation

Edge-finding algorithm for cell nuclei segmentation with ScanR analysis software.

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• Bacterial infection assay• Cell cycle analysis• Protein localisation and co-localisation• Live cell assays• Multicolour assays• Rare event analysis• Automated FISH analysis• Fluorescence analysis in tissue sections• Cell array screens• Micronuclei and comet assays

When is this technique the right choice?High-throughput fluorescence microscopy is the method of choice when automated and multi-parametric high-content analysis of many different samples or a large number of cells is needed.

What is required?Fluorescently labeled samples that would also be suitable for ob-servation with a normal fluorescence microscope. For example multi-well plates with transparent bottom, coverslips, etc.

How to get started?Contact VTT Medical Biotechnology for guidance and assis-tance.

Automated fluorescence (TurboGFP) intensity quantification and cell cycle analysis of siRNA transfected cells on Cell Spot Microarray.

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Availability within Turku BioimagingOlympus inverted microscope IX81 with ScanR

• VTT Turku.

ContactRami Mäkelä & Jeroen Powels, scanr@bioimaging.fi

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14. Stereo microscopy with microinjection

Stereo microscopy permits low-magnification study of samples. The long working distance and the availability of three-dimensional vi-sion make it possible to perform detailed operations under the mi-croscope. Combined with fluorescence and microinjection options, stereo microscopy becomes an all-round instrument for imaging purposes.

How does it work?In stereo microscopy, the light reflected from a sample to the eyes takes two separate optical paths. This produces a difference in viewing angle between the eyes, in turn permitting three-dimen-sional visualization of the sample. This effect depends greatly on the structure (thickness, opacity, and contrast) of the sample.

The Carl Zeiss stereo microscope.

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Properties of the BioImaging instrumentThe Zeiss SteREO Lumar.V12 fluorescence stereo microscope is a high-end instrument for low-magnification microscopy. Our 0.8x and 1.2x long working distance objectives allow a research-er to perform (for example) dissection using objective lenses permitting a maximal magnification of 120x. The versatility of the instrument permits imaging of (for example) entire mouse brain slices or, alternatively, single fluorescent cells, using the same objective. The instrument is equipped with both b/w and color CCD cameras to visualize fluorescent and non-fluorescent samples, respectively. The instrument has all of the basic fluores-cence filter sets and image acquisition is controlled by AxioVi-sion software.

What can be studied with this technique?Cell cultures, stained tissues, animal tissue, plant tissue, whole plants, zebrafish (hatched or as embryos), and insects can be visualized and imaged. Most material can be imaged with or without fluorescence.

Microinjection with the aid of the stereo microscopeAn Eppendorf InjectMan NI2 semi-automatic microinjection device has been installed beside the stereo microscope allowing (for example) injection into zebrafish embryos. Any material of small grain size can be microinjected. The most frequently in-jected materials are DNA, proteins, tracers, and chemicals with or without attached dyes.

Availability within Turku BioImagingZeiss SteREO Lumar• BioCity, 5th floor, room 5149.

ContactMarkku Saari & Jouko Sandholm, CIC@bioimaging.fi

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15. Time-resolved fluorescence microscopy

Time-Resolved Fluorescence (TRF) microscopy is an option worth considering when objects to be imaged exhibit high autofluores-cence or when extreme sensitivity is desired in low-throughput cell screening. In TRF microscopy, pulsed light excites labels and a time-gated mechanical chopper is used to set a detection delay. During this delay, shorter-lifetime emission derived from back-ground fluorescence and inherent autofluorescence dies out and only emission from photoluminescent labels with long decay times is detected. The technique is optimal for use with labels producing delayed fluorescence or phosphorescence.

How does it work?The time-resolved imaging workstation consists of a widefield upright fluorescence microscope (Nikon Eclipse E600) equipped with a prototype attachment (PerkinElmer Life Sciences, Wallac Oy) enabling both traditional prompt and time-resolved fluo-rescence imaging. The attachment includes a xenon flash lamp (pulse length 1 µsec; pulse energy 0.1 J; frequency 500 Hz), a me-chanical chopper, timing electronics, and a cooled CCD camera. The setup also includes a mercury arc lamp for standard fluores-cence imaging.

Signifer microimager.

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What can be studied with this technique?TRF microscopy is used in conjunction with photoluminescent labels with long decay times (e.g., lanthanide chelates of Eu and Tb, or Pd- and Pt-porphyrins). Time-resolved detection of long lifetime labels reduces background signal from sample autoflu-orescence, which lowers the label detection limit. As TRF mi-croscopy can resolve fluorescence emission with respect to both wavelength and lifetime, multiparameter imaging is a principal application of the technique. The TRF microscopy system is widefield in nature, and resolution is thus limited to the cellu-lar level. In low magnification applications, imaging of a 96-well plate is possible.

When is this technique the right choice?This technique is valuable when a label has a decay time of over 80 µsec. Usually, europium chelates are employed, and the tech-nique permits sensitive detection of cell surface proteins, pro-teins in tissue sections, or troubleshooting of microwell plate assays. More complex applications include multiparameter stud-ies, conducted when it is imperative to extract the maximum amount of information from a single cell using three or more separate labels on different epitopes.

Availability within Turku BioImagingSignifer microimager• BioCity, 5th floor, staircase A.

TRF microscopy images from adherent endothelial cells. Fluorescence from FITC-labeled cells (A) and autofluorescence of unlabeled cells (B) may be noted in the prompt channel. Fluorescence from Europium-labeled cells (C) and autofluorescence of unlabeled cells (D) may be observed in the time- resolved channel.

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ContactTuomas Näreoja, TRFM@bioimaging.fi

The principle of time-resolved measurement showing the decay curve of a long-lifetime label. The cycle commences with an excitation pulse and is followed, after 2000 microseconds, by another pulse.

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16. Live cell imaging

Live cell microscopy constitutes a versatile group of techniques that facilitate the imaging of cells over their entire life span. Rather than taking endpoint snapshots, it is possible to follow dynamic processes connected to events ranging from cell division to cell death.

What is live cell imaging, and why use it?Live cell microscopy affords the opportunity to visualize cellular events in real time. The same cells or cell population can be fol-lowed for hours. Turku BioImaging offers several live imaging platforms ranging from TIRF (total internal reflection fluores-cence) and high-content widefield fluorescence microscopy to high-throughput confocal microscopy. The primary require-ment is the establishment of conditions (CO2 level, humidity, and temperature) appropriate for normal cell function. Aberrant conditions can also be simulated. Hypoxia imaging (to 1% [v/v] oxygen) and imaging at below-ambient temperatures are also possible.

What can be studied with this technique?Available live cell applications include multi-position phase contrast and brightfield imaging with fluorescence. Also, FRAP (fluorescence recovery after photobleaching), FLIP (fluorescence loss in photobleaching), photoactivation, photoswitching, hy-poxia imaging, and multi-label time-lapse imaging are available. Almost all commercial fluorescent probes can be imaged with our instruments.

When is this technique the right choice?If it is important to understand the kinetic behavior of a fluores-cent target protein, or if scratch wound healing is to be recorded, live cell imaging is appropriate. Also, cellular processes such as endocytosis, apoptosis, cell division, and translocation can be visualized over time.

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Availability within Turku BioImagingTurku BioImaging offers numerous instruments suitable for live cell imaging, that include the following. Responsible operators are listed for each device and can be contacted through the com-mon email address below.

Zeiss LSM510 META confocal microscope• Fluorescence multi-label and multi-position confocal time-

lapse microscopy. Hypoxia imaging is possible. Uses 35mm glass-bottom plates or small chambers. Autofocus is laser-based.

• BioCity, 5th floor, room 5149 / Markku Saari & Jouko Sand-holm.

Leica TCS SP5 MP multiphoton confocal system• Suitable especially for situations where large penetration

depth is required, such as imaging living cells deep within their native tissue environment. Also useful if tissue fixation and labeling is not possible.

• BioCity, floor -1, staircase A / Tibor Veres.

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Zeiss AxioVert 200M widefield fluorescence microscope• Multi-label and multi-position fluorescence and/or phase

contrast time-lapse imaging. Imaging is possible with a 24-well plate incubator or a 35mm plate holder. Autofocus is software-based.

• BioCity, 4th floor, room 4086 / Markku Saari & Jouko Sand-holm.

Leica TCS SP5 MATRIX (HCS A)• Multi-label and multi-position fluorescence time-lapse imag-

ing. Phase contrast imaging currently not possible. Transpar-ent temperature control box covers the whole microscope, while the CO2-cotroller can be used either for well plates or single 35mm dishes. Accepts multiwall plates (up to 384 wells) or 35mm dishes and small objective glass-sized chambers. Several laser and software-based autofocus methods.

• BioCity, 2nd floor, room 2209 / Jari Korhonen.

Olympus CellR TIRF• Allows cell membrane TIRF imaging simultaneously with

widefield fluorescence imaging.• Medisiina, 2nd floor, room C204 / Jorma Määttä.

Zeiss LSM510 with JPK NanoWizard II atomic force micro-scope• Multi-label fluorescence confocal time-lapse microscopy and/

or atomic force time-lapse microscopy. Both the confocal and the AFM have incubator chambers with temperature control. The medium has to be CO2 independent as the instrument has no CO2 incubation arrangement.

• BioCity, floor -1, staircase A / Pasi Kankaanpää.

Carl Zeiss Axiovert 200M with Yokogawa CSU22 spinning disk confocal imaging unit• Suitable especially for studying fast dynamic events.• VTT Turku / Antti Arjonen.

IncuCyteTM HD (phase contrast imaging) and IncuCyteTM FLR (fluorescence imaging)• Live-cell imaging systems for studies of cellular growth, mi-

gration and morphology.• VTT Turku / Johannes Virtanen.

ContactLM@bioimaging.fi

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17. Image analysis

Whatever the imaging method used, image acquisition is only the beginning of the project. Processing, visualizing, and quantitatively analyzing image data is crucial in generating meaningful results. Turku BioImaging offers unique assistance in this area, thanks to in-house software production.

How does it work?In collaboration with the University of Jyväskylä, and other part-ners, a large open-source software package termed BioImageXD has been created, for post-processing of multidimensional biomedical image data. Acquired images are simply opened in BioImageXD, which can directly read common microscopy file formats and import regular image files. Hundreds of tools are available, ranging from basic processing such as noise removal and contrast adjustment to advanced tools including segmenta-tion, and quantitative and statistical analyses. Command pipe-lines can easily be created and saved, and hundreds of files can be processed, in a single session, according to these instructions. BioImageXD also creates three-dimensional views and four-di-mensional movies.

What can be studied with this technique?The possibilities are endless. BioImageXD can be used to analyze individual image pixels, to explore co-localization or to make in-tensity measurements within regions of interest. Structures such as cells or organelles can be extracted from an image using any of numerous segmentation methods, and parameters such as size, shape, distribution, and amount, can be quantified. The move-ments of cells or other objects such as intracellular vesicles can be tracked. The software can also perform bleaching and focus drift compensation, deconvolution, and automatic background

BioImageXD.

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removal to enhance image quality. Images can be interactively visualized in three dimensions, and distances (for example) can be measured. High-quality images and videos can be created for presentations and publications.

When is this technique the right choice?Image analysis is needed nearly every time a bioimaging method is used because images are useless without proper post-process-ing. BioImageXD is recommended because it is free and easy to operate, and offers most of the necessary tools in a single pack-age. The software is also actively developed and maintained in

Segmentation of cell nuclei prior to counting, using BioImageXD. Also shown are motion tracks related to endocytosis, drawn over a side view of a cell.

BioImageXD can create animations, for example by defining a flight path for the “camera”, as shown on the left. Such animations can lead into a cell, as displayed on the right, where a surface view of a nucleus inside a living cell can be seen.

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Turku, which means that help and user support, as well as tai-lored analysis, are available. No other software solution offers equal flexibility and support within Turku BioImaging. BioIm-ageXD is especially suitable for processing of fluorescence-based images, but many package tools are applicable to all kinds of im-ages.

What is required?BioImageXD will run on any normal personal computer, but powerful workstations are recommended for more demanding calculations or if large amounts of data are to be processed. The software is available free of charge for PC, Mac, and Linux. A basic understanding of digital images and the processing thereof is required, and, in many instances, some user training is needed.

How to get started?BioImageXD can be downloaded and installed on any computer (see below) An overview of the user manual provided may be helpful, as may attendance at a course or a training session or-ganized every year in Finland.

Availability within Turku BioimagingThe BioImageXD software is freely available for download at www.bioimagexd.net, and user support is available through info@bioimagexd.net. The email address below is for all other / general issues related to image analysis.

ContactPasi Kankaanpää, analysis@bioimaging.fi

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18. Flow cytometry

Flow cytometry is a method for analyzing cell populations. The technique allows thousands of cells to be evaluated. Most fluoro-phores available for fluorescence microscopy can be used in flow cytometry. The most common applications are phenotyping of cells, the counting of cells transfected with fluorescent proteins, and cell cycle analysis.

How does it work?Cells in suspension are injected into the instrument, where they are accelerated and hydrodynamically focused into a straight line of flow. The flow crosses a laser beam, and every cell is in-dividually illuminated. Fluorophores attached to or inside a cell are excited and the emitted light is gathered by collection optics. The optical system changes emitted light into an electrical signal, which is sent to a computer. In this manner, the levels of different fluorophores in each cell are recorded.

What can be studied with this technique?The starting material for flow cytometry analysis must be a single cell suspension, and the cells must be of diameter 0.5–50 μm. Cells can be stained with fluorophore-tagged antibodies, or may express an internal fluorescent protein or any other fluorescently labeled material (e.g., a probe measuring free intracellular cal-cium level). Different fluorophores with varying excitation or emission wavelengths can be simultaneously measured, depend-ing on the lasers and detectors fitted to the instrument. Thus, the intensities of fluorophores in/on individual cells are obtained. The staining of DNA with propidium iodide is an easy method for measuring cell cycle progression and apoptosis.

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When is this technique the right choice?Flow cytometry is ideal for end-point analysis, where the behav-ior of a cell population is to be studied. A large number of cells in a sample can be evaluated in a short time, and the number of samples can be easily increased using a high throughput sampler, enabling sample injection from a 96- or 384-well plate. Howev-er, flow cytometry is not ideal when the source of the emission must be known. The instrument cannot distinguish whether light emanates from the nucleus, cytoplasm, or other organelles. Only the overall levels of fluorophores associated with the cell are detected.

What is required?The cells to be analyzed must be prepared as a single cell suspen-sion. Many samples, including peripheral blood mononuclear cells (PBMCs), confluent cell lines, or bacteria, can be injected directly. Adherent cells must be detached. Tissue must be ho-mogenized before analysis. Practically all commercial fluoro-phores can be measured with flow cytometry. However, if it is desired to combine several different fluorophores in a single sample, attention should be given to excitation and emission wavelengths to prevent any possible overlap.

How to get started?Experimenters with no previous experience of flow cytometry should contact a responsible operator to request a training ses-sion. Training will cover all necessary steps, and will include tips for sample preparation, hands-on training on the instrument, how to set up the instrument for particular samples, and assis-tance in correct data analysis. The goal is to train all users to in-

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dependently operate the instruments. However, questions will be answered and help with advanced experiments will be available.

Availability within Turku BioimagingBD FACScan• BioCity, 5th floor, room 5126.

BD FACSCalibur• BioCity, 5th floor, room 5126, and Mikro, 1st floor, room

“Kurssityösali”.

BD LSR II• BioCity, 5th floor, room 5126.

ContactPerttu Terho, flow@bioimaging.fi

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19. Cell sorting

Flow cytometric cell sorting is a method for enrichment of cell pop-ulations, affording very high purity. This is an expansion of flow cytometry analysis. In addition to regular analysis, between one and four cell subpopulations can be selected and sorted into tubes. These cells can be re-cultured, or used for PCR or as a source of microarray data.

How does it work?As in basic flow cytometry analysis (please see previous chap-ter), cells are accelerated into the flow, a laser beam excites every cell individually, and the emitted light is collected and analyzed. After laser excitation, the flow is broken into drops. Each drop contains only one cell. If the cell inside the drop meets sorting criteria, the drop is electrically charged. The drop goes through deflection plates with a high potential, and charged drops are di-verted from the main stream into a collection tube. In this man-ner, every cell is analyzed and a decision on sorting is made for each individual cell. This means that the sorted cells are of very high purity.

What can be studied with this technique?Every problem amenable to analysis by flow cytometry can be addressed with this technique. The flow speed is typically about 1,000 cells/s.

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When is this technique the right choice?Cell sorting is the best option if it is desired to enrich GFP-pos-itive cells (or cells labeled with some other fluorescent protein). Also, antibody-labeled cells are typical sorting targets. If propid-ium iodide is used, cells in different stages of the cell cycle can be distinguished. Single cell sorting directly to wells of a plate is also possible. However, if very large numbers of target cells are needed, sorting may take too long. In such cases, pre-purifica-tion using (for example) magnetic beads is recommended.

What is required?Cell sorting is performed as a service. Thus, the details of the process are not important. Simply contact the responsible opera-tor and arrange a sorting time. It is always better to analyze cells by regular flow cytometry before sorting, to estimate the propor-tion of target cells. This helps to determine the time needed for cell sorting.

How to get started?Contact the responsible operator.

Availability within Turku BioimagingBD FACSAria IIu cell sorter• The sorter has two lasers, 488 nm blue and 633 nm red.• BioCity, 5th floor, room 5126.

ContactPerttu Terho, flow@bioimaging.fi

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20. Flow cytometry data analysis

The analysis of flow cytometry data is often an even bigger task than the sample preparation and measurement. In most cases, the researchers prefer to make the analysis with their own computers. Turku BioImaging offers Flowing Software, a free tool for analysing the datasets created by flow cytometers. The software package has been developed at the Turku Centre for Biotechnology, and sug-gestions for improvements and new features are always welcome.

What can be studied with this technique?Flowing Software is a flow cytometry data analysis software, which is able to perform all basic tasks, such as visualizing the data with dot plots and histograms, gating and calculating sta-tistics. The data copied from the instruments can be opened and analyzed in a very similar manner to commercial software pack-ages. The true power of the software becomes clear when the number of data files increases. Flowing Software offers unique tools for easy automation of analysis paths and the collection of results. Everything is based on objects and arrows connecting them. By connecting objects the user is able to create an analy-sis path, which performs all the tasks needed, from visualization to data collection. The automation does not require any special computational skills, because everything is done via the clear graphical user interface.

When is this technique the right choice?Flowing Software is suitable for all flow cytometry users, from beginners to demanding experts. The basic user is able to per-form basic analyses very easily. If one can use the software pack-

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ages that control the cytometers (e.g. CellQuest Pro, FACSDiVa Software), one can also use Flowing Software. For users running their samples from a well-plate, Flowing Software offers user-friendly tools for data collection. Many of the tools are unique and are often not even found in commercial software packages. For users requiring more complex processing of the data, Flow-ing Software is an excellent choice, as the analysis path always shows with arrows what is happening and how objects are in-teracting with each other. For developers with ideas of novel analysis methods, Flowing Software is the only choice, as it is possible to contact the development team and present ideas. For the developers it is a straightforward process to create a new ob-ject that executes the desired analysis step. The new object can be seamlessly connected to other analysis tools. In an ideal situ-ation, the time frame from the idea to the working solution can be just a few hours.

What is required?Flowing Software runs under Windows XP/Vista/7 environ-ments. The minimum requirements are quite low, but if the user wants to analyze very big data files (having millions of events), a powerful computer is recommended.

How to get started?The installation package is freely available from the software home page (see below). Flowing Software web page offers an orientation video, which shows all the basic and some of the advanced features. After watching the video, the user should be able to control the software with its full capacity. The develop-ment team of Flowing Software also arranges training sessions and workshops in the Turku area and, upon request, elsewhere.

Availability within Turku BioimagingFlowing Software can be downloaded from the addresswww.flowingsoftware.com.

ContactPerttu Terho, perttu.terho@flowingsoftware.com

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21. Atomic force microscopy

Atomic Force Microscopy (AFM) is a mechanical microscopy tech-nique, where images from biological samples can be obtained at atomic resolution, and forces between individual molecules or cells can be quantitatively measured.

How does it work?An Atomic Force Microscope (AFM) is a mechanical micro-scope, as opposed to (for example) an optical microscope. The principle is similar to an old-fashioned record turntable: a sharp tip attached to a flexible spring-like cantilever mechanically senses the force between the tip and the sample. Such forces cause the cantilever to bend, and the deflection is measured us-ing a laser beam reflected from the cantilever. The tip is atomi-cally sharp, so, in principle, a force between two single atoms can be measured. In biological AFM, an optical microscope is often additionally employed to adjust the system and to choose the sample area to be studied. The AFM can be operated in liquid and under physiological conditions, making it possible to study living biological specimens such as cells. The technique is label-free, making sample preparation straightforward. In addition to high-resolution images (AFM imaging) quantitative force meas-urements (AFM force spectroscopy) are possible using the same device.

What can be studied with this technique?AFM imaging can be used to obtain high-resolution (even at the level of an atomic lattice) images of fixed samples such as cells or a layer of protein attached to an even surface. The resolution is lower with living samples, but still much better than that af-forded by optical microscopy. AFM force spectroscopy can be

An atomic force microscope on the stage of a confocal microscope.

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used to measure the force between the tip and the sample, and if, for example, the tip is coated with a ligand, the position of a receptor molecule on the cell surface can be determined (recog-nition mapping). In addition, the tip can be replaced by an entire cell attached to the cantilever, and the attachment of this cell to a substrate such as collagen can be measured (cell adhesion meas-urements). The tip can also be used to physically manipulate a sample, or to pull on a protein to study how the protein unfolds.

When is this technique the right choice?AFM imaging is suitable when the resolution of optical micros-copy is not sufficient, when a sample needs to be imaged un-der native physiological conditions, or when the type of sample preparation required for electron microscopy is not possible. AFM is primarily used to obtain surface topography images. Thus, AFM is of limited use in studying the intracellular features of intact cells. Acquisition of an AFM image takes several min-utes, or even tens of minutes. AFM force spectroscopy is a good choice when binding forces and dynamics between molecules such as receptors and ligands, or between cells and a substrate, need to be determined. AFM can also be used to manipulate objects with high precision, for example to study the effects of specific mechanical stimuli on cells.

What is required?AFM is simple in principle, but some practical training is re-quired because an AFM is more sensitive than, for example, an optical microscope. In principle, living biological samples can be studied directly, but sometimes fixation is recommended, at least in the initial stages of a project. Samples need to be prepared on cover slips/in containers that are suitable for AFM. If force measurements are required, additional tip preparation may be needed, for example prior to coating with a ligand.

The principle of atomic force microscopy.

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How to get started?The operators responsible for the AFMs should be contacted to discuss the planned experiments and to determine how samples should be prepared and what type of cantilever should be used. Help can be provided on tip-chemistry when functional tips are required. AFM is not particularly rapid. It is important to allow sufficient time for experiments.

Availability within Turku BioImagingTurku Bioimaging offers two biological AFM setups with unique possibilities. Responsible operators are listed for both devices and can be contacted through the common email address below. AFM is also available in the Laboratory of Electron Microscopy (please see chapter 22).

JPK Nanowizard II combined with a Carl Zeiss confocal mi-croscope• Simultaneous imaging with the optical microscope. Scan

range 100 x 100 x 15 µm. BioCell for biological samples under physiological conditions. All standard imaging modes, and a CellHesion module that extends the Z range to 100 µm and enables cell adhesion measurements with a cell attached to the cantilever.

• BioCity, floor -1, staircase A / Pasi Kankaanpää.

Agilent ILM 5500, combined with either a Carl Zeiss wide-field fluorescence microscope or a Leica confocal/STED mi-croscope• Simultaneous imaging with the optical microscopes. Scan

range 100 x 100 x 8 µm. All standard imaging modes and soft magnetic AC mode for soft samples and simultaneous topog-raphy and recognition imaging for functional topography mapping.

• BioCity, 5th floor, staircase A / Tuomas Näreoja.

ContactAFM@bioimaging.fi

Confocal and atomic force microscopy images of the same cell and an example of a force measurement curve recorded using an AFM.

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22. Electron microscopy

In electron microscopy an electron beam, instead of light, is used to visualize the sample, resulting in very high resolution. The Labora-tory of Electron Microscopy at the University of Turku offers facili-ties for electron microscopic research, featuring microscopes as well as specimen preparation instruments and trained scientific and technical experts.

How does it work?Electron microscopy is a microscopic imaging technique us-ing an electron beam, instead of light, to visualize the sample, thereby allowing much greater magnification. The interactions of electrons with a specimen are used to provide information on fine structure. In Transmission Electron Microscopy (TEM), the reactions of electrons transmitted through an ultrathin (60 nm) section of a specimen are imaged. In Scanning Electron Microscopy (SEM), an electron beam falls at a non-vertical an-gle on a specimen and an image is derived from secondary or backscattered electrons emitted from the specimen surface after

A Jeol JEM-1200EX scanning-transmission electron microscope with an EDS analyzer (TEMSCAN).

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the impact of the primary beam. The use of electrons allows de-tailed imaging of structure, and localization and identification of molecules and elements to a resolution below 1 nanometer (10-9 m). The Laboratory of Electron Microscopy at the Uni-versity of Turku offers a full service, including consultation in the planning of experiments, sample processing, microscopic examination and recording of images, and reporting and inter-pretation of results. The goal of the research service is to provide electron microscopic facilities for investigators within the medi-cal faculty and in part also for scientists from all over Finland. In addition to basic morphological transmission and scanning electron microscopy, the facility offers immunocytochemistry, cryo-ultramicrotomy, X-ray microanalysis, scanning tunneling microscopy, atomic force microscopy, and morphometry.

When is this technique the right choice?If detailed structural details at nanometer resolution are re-quired, micro- and nano-structural imaging combined with mo-lecular analysis is the method of choice, applicable to samples of tissue, cells, bacteria, viruses, nanoparticles, macromolecules, and other related materials. Samples must be fixed with an al-dehyde, and dried either with ethanol or at the critical point of carbon dioxide (for SEM), and embedded in epoxy or another resin (for TEM) before being cut into thin sections.

The facility offers the following methods and instrumentation: • Cutting of ultrathin sections• Immunocytochemical light and electron microscopy• Recording and documentation of images taken using virtual,

light, and fluorescence microscopes, transmission electron microscopes (TEMs) and scanning electron microscopes (SEMs)

• An atomic force microscope (AFM)• A scanning tunneling microscope (STM)• X-ray spectrometry for element analysis and localization • Crystal analysis using electron diffraction

Availability within Turku BioImagingJEM 1200EX with EDS analyser (TEMSCAN)• 120 kV scanning transmission electron microscope with the

NORAN SIX EDS-system. The X-ray detector is a NORWAR-window Si(Li) instrument with an energy resolution of 145 eV. All elements heavier than boron can be analyzed.

• Medisiina, 4th floor, room A408.

Philips EM410 (TEM)• Medisiina, 4th floor, room A425.

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Nanoscope II tunneling and atomic force microscope (STM / AFM)• A total of four measuring heads are available, two for the STM

and two for the AFM.• Medisiina, 5th floor, room A512.

JEM 100SX• Conventional 100 kV transmission electron microscope with

a side-entry specimen holder. Specimen tilting is possible, to create stereo pairs.

• Medisiina, 4th floor, room A420.

ContactMarkus Peurla, EM@bioimaging.fi

Transmission electron micrograph of a thin section showing a portion of tes-ticular Leydig cell cytoplasm with a sector of the nucleus in the bottom righ cor-ner. The typical organelles that can be seen are smooth endoplasmic reticulum, rough endoplasmic reticulum, polyribosomes, and mitochondria.

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23. PET imaging

Positron Emission Tomography (PET) is a non-invasive method for imaging human biochemical and physiological processes in vivo. Minute amounts of biologically active compounds labeled with positron-emitting radionuclides are administered to subjects. The temporal and spatial distribution of these tracers within the body are visualized and measured using PET.

How does it work?Positron emitting radionuclides emit positrons (β+), which are the positively charged anti-particles of electrons. An emitted positron travels a few millimeters in tissue before colliding with an electron. The interaction causes annihilation of both particles, and the contained energy is converted into two photons (gamma rays), which are emitted in opposite directions. A PET scanner consists of detectors encircling a subject during the scan. The PET device registers only the coincidental signals of the two op-posite photons.

Positron-emitting radionuclides are usually produced in cyclo-trons. Short-lived (T½= 2-110 min) radionuclides are then in-corporated into molecules of interest using sophisticated radio-chemical syntheses. Because of the short half-lives of positron emitters, the syntheses must be very rapid. Tracers are delivered to the subject, usually by injection, and a PET scan is performed. Images are reconstructed as tomographic images by mathemati-cal processing of data.

What can be studied with this technique?• Tissue function and metabolism.• The functions of neurotransmitters and receptors.• Drug pharmacokinetics and drug dynamics.• Gene expression.

The annihilation process.

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When is this technique used?• PET is a versatile mode of molecular imaging (in vitro; ex vivo;

in vivo).• The very high sensitivity allows tomographic imaging of tar-

gets that are of low density.• The tracers do not have any pharmacological effects.• PET can easily be combined with anatomical imaging modali-

ties such as CT.• PET can be used in humans.

When is this technique the right choice?• PET is optimal when detection and/or monitoring of bio-

chemical or physiological processes in vivo is required.

What is required?• A suitable tracer.• Time slots for radiosynthesis and PET scans (projects are

scheduled four times yearly).• Expertise in the use of the technique, preferably by collaborat-

ing with existing experienced groups.

Availability within Turku BioImagingThe following instrumentation is available for preclinical imag-ing: small animal PET scanner, small animal PET/CT scanner, small animal ultrasound scanners, 4 human scanners for large animals. BioCity, floor -1, staircase B (first door to the left). Those interested are advised to contact the personnel at the Turku PET Centre preclinical imaging unit or at the TCDM unit, through the common email address below.

ContactPET@bioimaging.fi

PET methodology.

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24. Small animal ultrasound imaging

Ultrasound scanning is a technique used to visualize tissues and organs inside the body. It uses high-frequency sound waves to pro-duce images of different structures inside the body that can be dis-tinguished from each other by their different acoustic properties.

How does it work?When ultrasound waves are directed into the body, some are absorbed by body tissues and others bounce back. The sound waves that reflect back can be measured by the ultrasound ma-chine. Compared to many other imaging modalities, ultrasound is considered harmless and safe. Imaging by the high-frequency ultrasound device Vevo-2100 (by VisualSonics) is also called ultrasound biomicroscopy, referring to its almost revolutionary resolution of 30 microns. It can be used for visualizing most of the anatomical structures, ranging from a 5.5 days old mouse embryo to an adult rat. Compared to conventional ultrasound imaging, a transducer is attached to a probe holder and the ani-mal is anesthetized to optimize image quality. The platform is especially designed for evaluating hemodynamics in vivo in a non-invasive manner. Results are reproducible and reliable data obtainable for large cohorts being studied longitudinally. The device comes with an integrated micromanipulator allowing e.g. ultrasound guided injections for gene delivery or stem cell therapy via intramyocardial injections. Images and data sheets can be easily exported in multiple formats.

M-mode echocardiography from a 10 week old mouse (25g).

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What can be studied with this technique?The method can be applied on a large scale to translational ani-mal studies from cardiovascular research, cancer biology, devel-opmental biology and ophthalmology to reproductive biology and beyond.

When is this technique the right choice?Both the standard image resolution and the temporal resolution are very good, allowing visualization of small objects with cyclic deformation such as a mouse heart. Suitability for serial studies makes the method optimal for determining changes in develop-mental processes or disease progression.

What is required?Knowledge on how to apply this method for the research at hand is needed. Depending on the imaging protocol, for instance an ultrasound contrast agent for assessing tumor blood flow or specific software for acquiring three-dimensional data may be required.

How to get started?Only an “introductory lesson” is needed to get started with the method. Please contact the responsible operator.

Availability within Turku BioimagingVisualSonics Vevo-2100• Biocity, floor -1, room B131.

ContactJuha Koskenvuo, ultrasound@bioimaging.fi

B-mode echocardiography at diastole and systole (cut from live loop) from mouse with experimental apical infarct due to ligation of LAD coronary artery. White area visible both in epicardium and endocardium at anterior wall is the ligation stitch. Y-axis scale on millimeters.

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25. Micro-CT imaging

Micro-Computed Tomography (Micro-CT) is a method whereby X-ray computed tomography affords resolution on a micrometer scale. Micro-CT images can be used for volumetric analysis of scanned samples and three-dimensional image reconstruction.

How does it work?In micro-CT, hundreds of conventional X-ray images are ob-tained from a single sample, but with slight variation of the rota-tional angle. The spatial location of every voxel (a three-dimen-sional pixel) can be geometrically calculated from the images. Resolution of such images is, at best, on the micrometer scale, depending on sample size. The smaller the sample, the better the resolution.

What can be studied with this technique?Micro-CT is strictly an ex vivo technique. Micro-CT is princi-pally used for analysis of bone samples as part of basic research in bone biology, and in the study of various implant materials de-veloped to correct defects in bone. In addition, micro-CT can be applied to soft tissues, which, naturally, are of varying densities. Vasculature can be visualized after perfusion of a radiocontrast agent after sacrifice of the animal.

The use of micro-CT is somewhat limited because of the need for small sample sizes (maximally that of a thumb tip), and the rela-tively long times required for sample scanning (at least 45 min) and reconstruction (approximately 20 min/sample).

Visualization of otoliths in the balance organ of the mouse.

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Availability within Turku BioImagingTBI Skyscan 1072• Medisiina, room C204.

ContactJorma Määttä, CT@bioimaging.fi

Micro-CT visualization of the mouse proximal tibia.

A paraffin-embedded thorax of a rat pup, showing the chambers of the heart. Left: conventional X-ray image. Right: image reconstruction derived from the same sample.

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26. Imaging mass spectrometry

Label free imaging of tissues by Mass Spectrometry (MS) uses an intrinsic molecular parameter, the molecular mass, to determine and visualize the distribution of a wide variety of biomolecules at and on surfaces. The emerging field of research is known as Imag-ing MS (IMS).

How does it work?MS instruments function by measuring ionized molecules, where all values determined by MS are relative to the mass-to-charge ratio (m/z) assumed by the molecule after the ionization process, instead of the intact molecular weight. IMS uses an ionization technique termed MALDI that involves depositing a matrix (an aromatic organic molecule) on to a tissue which then seeps into the tissue. Irradiation of the matrix crystals leads to production of gas phase ions, stemming from an assortment of biomolecules (peptides, proteins, metabolites, lipids). The molecules are subsequently measured in a TOF (time-of-flight) mass analyzer. In the TOF ions are accelerated at a fixed voltage through a flight tube, where they become separated in time based on their m/z ratio, providing the ability to distinguish between biomolecules of different mass and to simultaneously measure their mass. Each position on the tissue section generates a mass spectrum describing the biomolecular content at that position. The mass spectra of an array of positions across the tissue section describe the biomolecular content of the tissue section and the spatial variation of every detected biomolecule within the tissue.

What can be studied with this technique?IMS allows the direct analysis and determination of the distribu-tion of up to hundreds of peptides and proteins directly from a

Imaging MS of rat brain. Mass peaks selected and visualized by color reveal the regional distribution and abundance of peptides. Combining the three individ-ual masses creates a composite image. In the overall mass spectrum plot, x-axis corresponds to mass-to-charge ratio (m/z) and y-axis denotes relative intensity.

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tissue section or tissue array. As there are no labeling techniques required molecules such as proteins, peptides, lipids, and me-tabolites can be analyzed.

Availability within Turku BioImagingIMS is conducted in the Turku Proteomics Facility, BioCity, at the Turku Centre for Biotechnology.

ContactGarry Corthals, MS@bioimaging.fi

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The IVIS Lumina II provides quantitative fluorescent and biolu-minescent imaging data from mice or rats in vivo. This is an ex-pandable, sensitive imaging system that is easy to use. The setup includes a highly sensitive CCD camera and an imaging chamber. Full automation is provided and the required analytic software is inbuilt.

How does it work?The system employs a light-tight imaging chamber. Inside the chamber, a heated stage is available for animals and anesthesia masks are built-in. The system has a CCD camera thermoelectri-cally (Peltier system) cooled to -90oC ensuring a low dark cur-rent and very little noise. The camera has high quantum efficien-cy over the entire visible-to-near-infrared spectrum. The stage is height-adjustable, allowing for a field of view of 5-12.5 cm. LED illumination is provided for reference photography, and a halo-gen lamp is available for fluorescence imaging. The system has a 10-position excitation filter wheel, and replaceable filter wheels to handle emissions at various wavelengths. All of these refine-ments are motor-controlled via IVIS Living Image software.

27. Macroscopic in vivo imaging

Mice with orthotopically inoculated PC3M-prostate cancer cells carrying a luciferase reporter imaged with IVIS Lumina II.

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What can be studied with this technique?Cells and particles expressing luciferase or other fluorescent molecules can be visualized with the IVIS Lumina II system. Macroscopic in vivo imaging enables (for example) phenotyping of transgenic mice, imaging of growing tumor cells in a living animal, follow-up of treatment responses, visualization of in-flammation and infections, and arrival of nanoparticles at des-ignated targets. A principal application area is the study of on-cological xenografts, in which cancer cells with bioluminescent/fluorescent reporters are inoculated into immunodeficient mice/rats. These oncology models can be used to assess anti-cancer therapies over the course of treatment in vivo. Non-invasive im-aging of tumor growth and metastasis allows longitudinal evalu-ation of tumor development before, during, and after treatment, offering an excellent preclinical strategy for assessment of tumor response and recurrence.

When is this technique the right choice?IVIS Lumina is appropriate when fast or repeated imaging is re-quired and a large number of animals is to be screened.

Availability within Turku BioImagingIVIS Lumina• BioCity, floor -1, room YK136.

ContactMarko Tirri, IVIS@bioimaging.fi

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Laser-capture microdissection is a technique used to isolate cell(s) from a certain area to analyze their properties. This isolation method yields pure cell populations. Thus, neighboring, unwanted, different cells do not bias the results.

How does it work?Laser-capture microsdissection (LCM) technology uses focused laser beam to extract the material of interest. The Zeiss PALM microdissection device employs a focused beam to cut the de-sired material, and a defocused beam to catapult that tissue into the cap of a tube. The catapulting process is upward, thus pre-venting the entry (by gravity) of unwanted material into the cap. The source material can be placed on a normal objective glass slide or on a membrane slide. In the latter case, the desired mate-rial is excised and catapulted together with a piece of membrane.

What can be studied with this technique?The system can be used with frozen or paraffin-embedded tissue samples, single cells, and even single chromosomes. The micro-scope is equipped with a fluorescence system and has a video camera to assist in target visualization. In PALM, separation of the specimen is performed with a laser beam focused so precise-ly that a beam accuracy of less than 1 µm can be achieved. Avail-able objectives are the FLUAR 10x/0.50 and LD ACHROPLAN 40x/0.60 for dissection, and the LD ACHROPLAN 20x/0.40 for specimen visualization.

When is this technique the right choice?Microdissection is appropriate when a distinct cell population is to be isolated from a sample. Such a population could (for exam-

28. Laser-capture microdissection

Carl Zeiss PALM.

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ple) be a group of cancerous cells, a particular part of an organ, green fluorescent cells, or fast-response muscle cells. Different populations can be chosen from the same sample.

What is required?For successful downstream processing, the sample must be treat-ed prior to LCM with an eye toward the downstream analysis that is planned. A good-quality sample is essential for satisfac-tory results. Downstream testing may include (for example) mi-croarray analysis, PCR, quantitative RT-PCR, and/or protein/enzyme detection.

Availability within Turku BioImagingZeiss PALM• BioCity, 5th floor, room 5149.

ContactMarkku Saari & Jouko Sandholm, CIC@bioimaging.fi

Left: uncut tissue. Right: tissue after microdissection.

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An international MSc Degree Programme in Biomedical Imaging commenced in Turku in September 2010. The programme is jointly administered by the Faculty of Medicine at the University of Turku and the Department of Biosciences at Åbo Akademi University.

Structure of the programmeThe interdisciplinary curriculum consists of major subject stud-ies (90 ECTS) and optional studies including advanced courses in microscopy, photonics, in vivo imaging, image analysis, and multimedia editing, as well as an international lecture series and hands-on courses. As imaging is a practical science, the pro-gramme provides students with good practical skills in imaging methods ranging from molecular and cellular to tissue and clini-cal imaging.• Duration: Two academic years (120 ECTS).• Acquired degree: Master of Science (MSc).• Language of instruction: English.• Possibility of participation in high-end local research in dif-

ferent application areas or of an exchange with international university partners to fulfill Master’s thesis work.

Who may apply?The programme is intended for students with a BSc degree in the Life Sciences or relevant areas of biomedical sciences, physics, chemistry, or engineering. The applicants should also be familiar with the basics of laboratory practice. Both Finnish and foreign students are welcome to apply for the international programme. The annual intake of the programme is a maximum of 30 stu-dents.

29. Master’s Degree Programme in Biomedical Imaging

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Language requirementsA good knowledge of the English language is required. All ap-plicants must prove that they have adequate English skills by ap-pending the score from an international English-language test to their applications.

Aims of the programmeBiomedical imaging has emerged as one of the most important technologies in basic bioscience and biomedical research, as well as in clinical medicine and translational studies. The cur-rent technologies enable detailed real-time non-invasive visuali-zation of molecules, structures, and events in cells, tissues, and whole organisms. Application areas include cellular and molecu-lar biology, pharmacology, structural biology, nanotechnology and biomaterials research, and patient care and diagnostics. The Programme in Biomedical Imaging aims to give professionals a thorough understanding of diverse imaging technologies along with practical skills. The advanced portion of the curriculum is also intended to offer training to graduate students and post-doctoral fellows.

CollaboratorsThe immediate collaborators are the national Turku PET Centre, the Turku Centre for Biotechnology, the Turku Center for Dis-ease Modeling (TCDM), and the Turku University Hospital. The programme has collaborators in the Nordic countries, Europe, the USA, and Asia, and also offers courses given by international experts.

Career prospectsThe interdisciplinary curriculum provides graduates with an ex-cellent basis for careers in many different areas of life sciences, both in academic research and industry. Importantly, the pro-

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gramme will give students considerable latitude to tailor their curricula to their needs, depending on previous background and future career interests in the imaging field.

More information:www.med.utu.fi/biomagingwww.abo.fi/bioimaging

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30. Turku as an imaging stronghold

Turku has a genuine tradition in imaging research. Over the past few years, Turku has established outstanding competence in a broad array of imaging areas. Many bioimaging initiatives are collaborative efforts between the University of Turku and Åbo Akademi University. Examples of very successful joint facilities are the National PET Centre and the well-established Cell Imag-ing Core (CIC) at the Turku Centre for Biotechnology.

Many Departments in the University of Turku Medical Faculty have a tradition in different areas of imaging. An example is the Laboratory of Electron Microscopy in the Institute of Microbi-ology and Pathology. Recently, a National Center for Disease Modeling has been established in Turku. This will be a significant asset for many groups working on imaging and will in turn ben-efit from the availability of the advanced imaging infrastructure required to visualize different types of disease processes.

The Turku bioscience unit of the State Technical Research Cen-tre (VTT) has placed strong emphasis on imaging-based high throughput screening approaches. Furthermore, the Turku Cen-tre for Biotechnology hosts the national Microarray Centre, to-gether with the CIC, bringing genomic imaging within the reach of many research groups in Turku and elsewhere. The Centre for Biotechnology also has a rapidly growing core unit in proteom-ics, including mass spectrometry-based imaging of the proteome and post-translational modifications thereof.

The Chair of Turku BioImaging, Professor John Eriksson (left), and researcher Kimmo Isoniemi (right), explain new three-dimensional imaging technology to the Rector of Åbo Akademi, Jorma Mattinen, and the Rector of the University of Turku, Keijo Virtanen.

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It should also be noted that, beginning in the early 1990s, Turku has become one of the world’s leading cities in the development of new optical technologies for microscopy and bioanalytics. Some research results have been successfully commercialized.

Turku’s strengths include imaging-related research in physics and chemistry and the presence of a community active in in-formation science, all of which give a competitive edge to local imaging. Within the field of information science and process technology, there is growing interest in imaging technologies, systems biology, and modeling of cellular processes. Bioimaging thus bridges seamlessly with the engineering sciences.

Finally, many different enterprises associated with imaging-ori-ented research and product development have facilities in Turku. These include Perkin-Elmer Wallac, Turku Imanet, Arctic Diag-nostics, and several other companies and affiliated contractors. The city thus has a tradition of commercial production of tech-nologies relevant to imaging.

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Turku BioImaging email addresses

Turku BioImaging recommends using these email addresses to contact the people responsible for the various techniques pre-sented in this booklet. First part of the address is the bolded part below, followed by @bioimaging.fi. Each address can also be found in the chapter introducing the corresponding technique.

LM = all light microscopy techniquesCIC = cell imaging coreconfocal = confocal microscopyHT = high-throughput microscopySTED = high-resolution microscopyFRET = fluorescence resonance energy transfer microscopyFCS = fluorescence correlation spectroscopy & raster image correlation spectroscopy2-photon = two-photon microscopyspinning = spinning disk confocal microscopyTIRF = total internal reflection microscopyscanr = scanR automated fluorescence microscopeTRFM = time-resolved fluorescence microscopyanalysis = image analysis & processingflow = flow cytometry & cell sortingAFM = atomic force microscopyEM = electron microscopyPET = positron emission tomographyultrasound = ultrasound imagingCT = micro-CT (computed tomography) MS = mass spectrometry imagingIVIS = macroscopic in vivo imaging

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EditorsJohn ErikssonPasi KankaanpääJari KorhonenMaritta LöytömäkiEeva RainioJouko Sandholm

Image editorsPasi KankaanpääMaritta LöytömäkiMarkku Saari

DesignJoacim Päivärinne

Printed byFinepress OyOctober 2011

Authors

1. What is Turku BioImagingText: Maritta Löytömäki & John ErikssonImage: Sami Koho

2. A Brief history of Turku BioImagingText: Maritta Löytömäki & John ErikssonImages: Markku Saari / Turun yliopiston viestintä

3. Turku BioImaging facilitiesText: Maritta Löytömäki & John ErikssonImages: Diana Toivola / Markku Saari / Heikki Minn / Mika Okko / Lauri J. Pelliniemi / Pasi Kankaanpää

4. Light microscopyText & image: Jouko Sandholm & Markku Saari

5. Confocal microscopyText & images: Pasi Kankaanpää

6. High content analysisText: Kimmo Isoniemi & Jari KorhonenImages: Kimmo Isoniemi / Vesa-Matti Väärä

7. STED microscopyText & images: Sami Koho

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8. FRET microscopyText & image: Daniel Abankwa

9. FCS and RICSText & images: Daniel Abankwa & Camilo Guzmán

10. Two-photon microscopyText & images: Tibor Veres

11. Spinning disk confocal microscopyText: Antti ArjonenImages: Yokogawa Electric Corporation / Antti Arjonen

12. TIRF microscopyText: Jorma MäättäImage: Antti Arjonen

13. ScanR automated fluorescence microscopyText: Rami Mäkelä & Jeroen PowelsImages: Juha Rantala

14. Stereo microscopy with microinjection Text: Jouko SandholmImage: Markku Saari

15. Time-resolved fluorescence microscopyText: Tuomas Näreoja & Roope HuttunenImages: Pasi Kankaanpää / Roope Huttunen / Tuomas Näreoja

16. Live cell imagingText: Jouko Sandholm & Markku SaariImages: Pasi Kankaanpää / Markku Saari

17. Image analysisText & images: Pasi Kankaanpää

18. Flow cytometryText: Perttu TerhoImages: Perttu Terho / Perttu Terho / Markku Saari

19. Cell sortingText & images: Perttu Terho

20. Flow cytometry data analysisText & image: Perttu Terho

21. Atomic force microscopyText & images: Pasi Kankaanpää

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22. Electron microscopyText: Lauri J. PelliniemiImages: Björn Enberg & Lauri J. Pelliniemi / Lauri J. Pelliniemi

23. PET imagingText: Tove GrönroosImages: modified from Cherry & Gambhir (Ilar J. 2001) / Tove Grönroos

24. Small animal ultrasound imagingText & images: Juha Koskenvuo

25. Micro-CT imagingText & images: Jorma Määttä

26. Imaging mass spectrometryText: Garry CorthalsImage: Olli Kannaste

27. Macroscopic in vivo imagingText & image: Marko Tirri

28. Laser-capture microdissectionText: Jouko SandholmImages: Markku Saari / Jouko Sandholm

29. Master’s Degree Programme in Biomedical ImagingText: Maritta Löytömäki & John ErikssonImages: Iloinen Liftari / Pasi Kankaanpää / Pasi Kankaanpää

30. Turku as an imaging strongholdText: Maritta Löytömäki & John ErikssonImage: Michael Karlsson

Notes

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