PD Dr. Bernhard Gaese

Contents

The practical teaches techniques to determine auditory function and dysfunction in rodents. These techniques can be used to determine effects of pharmacological or behavioural treatments of sensory disorders such as tinnitus or hearing loss. The focus is on behavioural techniques suitable to characterize the disorder rather precisely in comparison to normal functions. All steps that are necessary for a project in the field are taught in this practical: study design, animal handling, control of experimental parameters, pharmacological treatment of animals, and data analysis. The behavioural analysis is paralleled by basic electrophysiological measurements necessary to determine the effects of dysfunction and treatments at the physiological level. The students work on their own projects under supervision and present their results in the form of a seminar talk. The main focuses are: measuring and analysing behavioural data, performing efficient physiological experiments to determine auditory function, and statistical evaluation methods. Preparation of a potential publication will be the final part of the project. After completion, the individual projects will be presented and discussed in the form of a seminar talk. In a further seminar talk the students will present an original piece of research from the area of cognition and hearing.

Educational objectives / Competences

Familiarity with carrying out well controlled behavioural experiments (animal handling, measuring and analysing behavioural data, statistical analysis). Performing physiological measurements including electrophysiological recording in minimally invasive preparations. Additional aspects are: introduction to software for data handling, signal processing, and graphical display. Deriving scientific questions from the current literature. Knowledge about the usage and limitations of animal models for neurological diseases.

PD Dr. Bernhard Gaese

Contents

The practical covers the whole range of techniques to investigate brain activity underlying the processing of sensory information in the auditory domain. The focus is on electrophysiological single cell techniques in rodents in the awake and anesthetized preparations. Brain activity is acquired and analysed with the goal to understand behavioural responses following auditory stimulation. Cognitive aspects (e.g. context-dependence) are taken into account. The students work on their own projects under supervision and present their results in the form of a seminar talk. The main focuses are measuring and analysing neuronal activity in different configurations of in-vivo recording techniques. The following analysis includes modern techniques of signal processing, efficient handling of larger data sets and statistical evaluation methods. Preparation of a potential publication will be the final part of the project. After completion, the individual projects will be presented and discussed in the form of a seminar talk. In a further seminar talk the students will present an original piece of research from the area of cognition and hearing.

Educational objectives / Competences

Familiarity with carrying out physiological experiments (animal handling, surgery, measuring and analyzing electrical activity at the single neuron level. Combining physiology with neuroanatomical and histological staining techniques. Basic introduction to behavioural control. Introduction to software for data handling, signal processing, statistical analysis and graphical display. Understanding cognitive influences on sensory information processing as an important aspect of context-dependent behaviour. Deriving scientific questions from the current literature.

Prof. Dr. Bernd Gruenewald

Contents

In the practical course, the physiological foundations of behavior control are examined. Students work on their own projects, the topics of which have been jointly defined in advance. The techniques taught include: Cell physiology (patch-clamp recordings, intracellular recordings, calcium imaging, cell culture); neuroanatomy (staining methods, brain preparations, confocal laser scanning microscopy, fluorescence microscopy); behavioral experiments (behavioral pharmacology, extracellular recordings, learning and memory, social behavior). Insects (honeybees, Drosophila) are used as model organisms. The main topics are: Functioning of ion channels and transmitter receptors, neuromodulation, learning behavior, olfactory memory formation, social behavior of honeybees. The student presents his or her results in the form of a seminar presentation and a poster. In a further seminar presentation, original physiological and behavior-analytical papers are critically reviewed. The presentations are given in English and the student receives detailed feedback on the content and form of the presentations. By writing a protocol in the form of a paper, students familiarize themselves with writing a scientific publication. From planning to conducting, recording and evaluating the original data, the student essentially works independently once the individual work steps have been taught.

Educational objectives / Competences

 The student plans behavioral-physiological experiments, carries them out and evaluates them. Knowledge of the measurement of ion currents, behavioral observations and behavioral quantification as well as neuroanatomical methods is acquired during the module. The student learns how to approach scientific questions and literature studies.
The preparation of scientific papers and giving presentations are taught and learned.

Prof. Dr. Ernst H.K. Stelzer & Dr. Francesco Pampaloni

Contents

This practical course teaches the basics of three-dimensional cell cultures and modern three-dimensional microscopy. A significant development in recent years is the return to the observation of living biological samples in a physiologically relevant context. Cells are cultivated and examined under physiological conditions. These conditions are ensured in tissue pieces and three-dimensional cell cultures in collagen or in other tissue-like hydrogels of the extracellular matrix (ECM), such as Matrigel. The quantitative analysis of living three-dimensional structures requires rapid optical sectioning. Confocal fluorescence microscopy is only suitable for relatively thin samples, as the signal is rejected by the pinhole aperture of large, highly scattering objects. The energy efficiency (ratio between the energy that excites the specimen and the energy that is detected) is therefore low. One possible approach is the consistent application of light sheet-based fluorescence microscopy (LSFM) in combination with the three-dimensional preparation of specimen, which represents an overall concept. The students work on current research projects of the Stelzer group under supervision and present the results in the form of a protocol (report) and a seminar presentation.

Educational objectives / Competences

The student masters the basic concepts of classical two-dimensional and three-dimensional cell culture. He or she will describe various applications of three-dimensional cell cultures and usable cells in the life sciences. She or he is familiar with the fundamentals and basic concepts of classical microscopy (properties of light, resolution, aperture) and photometry (energy, power). She or he is familiar with the differences between confocal and light sheet-based fluorescence microscopy and knows the limits of classical microscopy in dense tissues. She or he has the skills to prepare, isolate and stain spheroids, cysts, organoids and three-dimensional tissue sections. She or he prepares samples for the various microscopes, records image data sets, processes and evaluates the data.

Prof. Dr. Enrico Schleiff & Dr. Sortirios Fragkostefanakis

Contents

The practical course teaches basic working techniques and experimental concepts of molecular cell biology in general and specifically on questions of cellular and molecular plant physiology. Main focuses are: protein biochemical methods for the study of protein translocation and chloroplast dynamics, including subcellular fractionation, basics of working with plant cell cultures and transgenic plants, in vivo and in situ measurements of activity and localization, including digital image processing. In addition, students investigate the response of plant cells to conditions that cause proteotoxicity and how cells use protective mechanisms to survive and recover from stress. The focus is on the regulation of the capacity and activity of chaperones. Students learn how to work with transgenic plants, eukaryotic cell cultures and protoplasts, i.e. their cultivation, passaging and transfection for ectopic expression or protein silencing. The analysis includes a broad spectrum of molecular biological and cell biological techniques such as PCR, cloning, SDS polyacrylamide gel electrophoresis and western blotting, immunofluorescence, protein activity measurements, etc. The students work on current projects under supervision and present the results in the form of a seminar presentation. In a further seminar, they present an original work from the field of cellular and molecular plant physiology. They learn how to write a scientific paper by preparing a protocol of the results (report).

Educational objectives / Competences

The student acquires knowledge of the isolation of plant cell organelles and the independent characterization of organelle proteins and masters the handling of sterile work and the cultivation and transfection of cells. In addition, the student works independently on the fluorescence microscope and applies the computer-aided evaluation of laboratory data and image files. Knowledge of the analysis of transgenic plants as well as independent processing of scientific questions against the background of relevant literature will be acquired.

Prof. Dr. Ingo Ebersberger

Contents

This practical course teaches basic methods and algorithms for the bioinformatic analysis of large sequence data sets. Considering current data from high-throughput sequencing, students work on questions relating to the functional characterization and evolution of physiological metabolic pathways and protein complexes. The focus is on the preparation of new sequence data sets for analysis, data mining to complement existing data sets and bioinformatic methods for the comparison and annotation of sequences.  The theoretical basis of these analyses is formed by independent literature work and a seminar presentation on an original work from the field of applied bioinformatics. By summarizing the results at the end of the practical course in a seminar presentation and in writing in the form of a report, students learn how to present scientific research results correctly.

Educational objectives / Competences

The student performs functional sequence annotation, bioinformatic annotation transfers and the prediction of functionally equivalent proteins, taking evolutionary relationships into account. The student deals with large sequence data sets and analyzes them bioinformatically, mines public databases and describes relational database systems. The student creates and interprets phylogenetic profiles and masters the basics of independent processing of scientific questions against the background of relevant literature.

Prof. Dr. Zoe Waibler

Contents

During the practical course, students work on the current projects of the working group. The focus is on immunological experiments with primary human cell cultures, occasionally also in the murine system. The analyses cover a broad spectrum of immunological and cell culture techniques such as FACS, ELISA, plaque assay, viral infections, (q)RT-PCR, the isolation of different cell types from human blood donations and the separation of cells using MACS and cell sorters. The results of the practical course are presented by each student in the form of a written protocol and a presentation at the end of the course. The students also participate in the weekly lab meetings where they are informed about the ongoing research projects of the group and report themselves.

Educational objectives / Competences

Students plan and carry out complex immunological experiments. Students critically evaluate current literature in the field of immunology.

Prof. Dr. Didier Stainier

Contents

Theoretical and experimental principles of developmental biology and genetics are taught in the practical course. Research focuses on the development, function and regeneration of the cardiovascular system. Another research focus of the lab is a newly identified process called transcriptional adaptation, in which mRNA degradation products modulate gene expression. Students work on current research projects and investigate cellular and molecular processes relevant to the above research areas. Experimental work includes genetic testing methods in a range of model systems including zebrafish, mouse, C. elegans, Neurospora and mammalian cells in culture.  Performance of live imaging of zebrafish embryos and larvae, preparation of tissues for in situ hybridization, immunohistochemistry, immunofluorescence microscopy, confocal microscopy; performance of molecular biology techniques and handling of zebrafish and other models (e.g. DNA and RNA injections into zebrafish embryos).  The experiments of the practical course are summarized and recorded in writing by the students and presented at the end of the course.  The students participate in the weekly seminars of the working group, where they are informed about current research topics of the working group.  In literature clubs, students are familiarized with current scientific publications on the topic of the practical course.

Educational objectives / Communication

Students master the basic techniques of molecular biology and genetics and are familiar with the use of model systems. They practice working with original literature in English.  Students acquire the competence to present and discuss experimental results in an international environment.

PD Dr. Boris Strilic

Contents

The practical course provides basic knowledge and various working techniques in the field of general cell and molecular biology and, in particular, endothelial and tumor cell biology. Students work on their own projects under supervision, which are then carried out in the laboratory. Students analyze their data both quantitatively and qualitatively and present the results in the form of a written protocol (report). Students also take part in weekly seminars given by other lab members. Working with animal models under supervision is possible depending on the project.

Educational objectives / Competences

The practical course serves to learn various techniques from the above-mentioned areas. The student masters the cultivation of various eukaryotic cell lines and primary cells, siRNA-mediated knockdown, the preparation of histological sections with subsequent immunofluorescence staining and evaluation using a confocal laser microscope, (quantitative) PCR, Western blots and immunoprecipitation. The student presents and discusses the results in an international environment.

Prof. Dr. Virginie Lecaudey

Contents

The practical course teaches the theoretical and experimental basics of cellular developmental biology. Research focuses on the mechanisms of cell migration and the morphogenesis of organ formation using the zebrafish model. Our main model system is a group of about 100 epithelial cells that migrate as a group over a long distance in the embryo. The students participate in the scientific experiments of the working group and investigate mechanisms of cell migration, cell differentiation, changes in cell morphology or cell proliferation in this system or animal model. Techniques used include zebrafish handling (crossing, injection, genotyping), genetics (generation of mutants and fluorescent transgenic lines), molecular biology and modern live imaging and image analysis techniques. The results of the practical course are presented by each student in the form of a written protocol. Students also participate in weekly lab meetings where they are informed about the ongoing research of the members of the group. In a literature seminar, each student presents a current publication on his or her project.

Educational objectives / competences

Students master the basic techniques of molecular and developmental biology, including the handling of zebrafish and modern live imaging techniques as described above. They work with a vertebrate model organism, present their data orally and comprehensibly and document their experiment in a lab book. The students are in an international environment and write and communicate their results in English.

Prof. Dr. Michaela Mueller-McNicoll

Contents

In the practical course, students learn to cultivate mammalian cell culture models and, with the help of CRISPR/Cas9 genome editing, to label endogenous human genes coding for RNA-binding proteins (RBPs) with a GFP marker (green fluorescent protein). The GFP marker makes it possible to visualize the tagged proteins in living and fixed cells using
fluorescence microscopy and to determine their subcellular localization or their location in different cellular sub-compartments, e.g. paraspeckles, nuclear speckles, stress bodies or stress granules. During the practical course, students will design and clone plasmid constructs for GFP labeling using Gibson assembly, transfect cells to enable genome editing, and verify successful GFP labeling by Western blot, genomic PCRs, RT-PCR, and confocal microscopy. Changes in the subcellular distribution of RBPs under stress will be monitored by live cell imaging, immunofluorescence microscopy and RNA fluorescence in situ hybridization (FISH). Other techniques include rapid depletion of RBPs, subcellular fractionation, differentiation of cells into neurons, various stress treatments, image analysis and quantitative analysis.

Educational objectives / Competences

Students organize and plan their experiments themselves. They carefully evaluate their results, document the data and quantify it. They work in a team and communicate their data to colleagues in English and write protocols (report). They critically evaluate and discuss scientific articles.

Prof. Dr. Manfred Koessl & Dr. Julio Hechavarria

Contents

The main goal of this course is to understand how mammals communicate using acoustic information (sounds). The course is designed from the perspective of the “broadcaster-receiver" approach, and therefore it is consequently subdivided into two parts. The first part is meant for understanding the sounds broadcasted by two mammalian species (Mongolian gerbils and bats) while they are communicating. Basically, using bioacoustics tools, the students will try to figure out the vocal alphabet of bats and gerbils. The second part of the course deals with the receiver. In this part, the students will learn how the gerbil's voice is processed in the brain by neurons located in the auditory cortex. The main aim here is to assess what happens in the brain when an animal hears a behaviourally relevant sound. At the beginning of each course part, there will be introductory discussions that will provide the students with the necessary theoretical background for conducting and understanding the different experiments. An introduction to statistics and to MATLAB will also be offered. The final report will be written in the form of a scientific paper, and the results will be presented in the form of a short talk.

Educational objectives / Competences

By the end of the course, the students should be able to: (1) Understand basic concepts of bioacoustics such as the sound as a mechanical wave, sound transduction using microphones, analogue-to-digital conversion using sound cards. (2) Measure basic parameters of a sound wave (frequency, duration, intensity). (3) Perform basic surgeries required for acquiring neuronal data. (4) Understand basic neuroscience concepts such as: action potential, local field potential, receptive field, brain topography,
spike clustering, brain oscillations. (5) Testing hypothesis using basic statistical tests (normality tests, parametric and non-parametric t-tests and analyses of variance (ANOVA)).

Prof. Dr. Amparo Acker-Palmer & Dr. Bettina Kirchmaier

Contents

The practical course provides basic theoretical and experimental knowledge in the field of cellular, molecular and systemic neurobiology in mice and zebrafish. Students work on their own projects under supervision and present the results in the form of a lecture. In a further lecture, they present an original paper from the topic area of their projects. They learn how to write a scientific paper by preparing a protocol (report) of the results. 
The practical course is divided into two units. The first part includes the following work: Basic mouse genetics techniques, processing brain tissue for immunohistochemistry, basics of working with cell cultures, including the preparation of primary neuronal, astrocytic or endothelial cultures; immunofluorescence microscopy, confocal microscopy and  biochemical techniques including protein gel electrophoresis and western blot. In the second part of the practical course, students are familiarized with basic genetic techniques in zebrafish research. This includes learning molecular biological and histological methods, the use of different microscopes, the manipulation of zebrafish embryos and the performance of simple behavioral tests.

Educational objectives / Competences

Students master the basic techniques of cellular, molecular and systemic neurobiology. They work independently and sterilely on cultured cells, use the fluorescence microscope and stereomicroscope independently, carry out basic zebrafish work such as handling embryos and genetic techniques, and perform computer-aided analysis of laboratory data and image files. Students work in an international environment and present and communicate their results in English.

Prof. Dr. Franziska Matthaeus

Contents

The module teaches the basics of theoretical biology based on a project on the dynamics of multicellular systems that students work on themselves. The working techniques used can be in the field of data or image analysis (segmentation, object identification, tracking, dimensional reduction of large data sets, motion analysis) or in the modeling and simulation of multicellular systems. Areas of application include collective phenomena, self-organization and “muter" formation (e.g. in development processes or experimental model systems). Alternatively, the data or image analysis, modeling and simulation can also relate to a previous project carried out during the Master's degree course, from which suitable data is available.
Some of the image analysis can be carried out using existing software (ImageJ, Matlab). For further processing of the data, visualization or statistical evaluations, own programs should be created in a known or to be learned programming language (e.g. Python, Matlab, Julia). The modeling and simulation also involve working on and developing software. The student presents the results of the project in the form of a graded report and a seminar presentation.

Educational objectives / Competences

Students have basic or in-depth programming skills. They are familiar with modeling approaches, data analysis techniques, image processing methods and can use and further develop image processing software and modeling approaches independently at a later stage.

Prof. Dr. Stefan Eimer

Contents

In this practical course, students will learn how to use the multicellular model organism C.elegans to investigate the early molecular mechanisms and cellular changes that lead to the development of Parkinson's disease. By inactivating and analyzing homologous genes that are associated with the hereditary form of Parkinson's disease, an attempt is made to understand the function of these genes in normal cells on the one hand, and on the other hand to investigate which cellular changes result from the inactivation of these genes, which allow conclusions to be drawn about possible early mechanisms of the development of Parkinson's disease. The mutant animals produced will be analyzed genetically, biochemically and cell biologically using high-resolution microscopy techniques such as confocal microscopy and electron microscopy.

Educational objectives / Competences

Students understand and recognize complex cellular relationships that can lead to the development of a neurodegenerative disease through genetic manipulation and analysis in a multicellular model organism. Participants will master the use of the C.elegans model system and state-of-the-art methods of genetic manipulation such as CRIPR/Cas9-mediated genome editing or RNAi gene knockdowns as well as the generation and quantitative analysis of microscopy data.

Prof. Dr. Jasmin Hefendehl

Contents

The practical course provides basic theoretical and experimental knowledge in the field of neurodegenerative and vascular diseases. Students work on their own projects under supervision and present the results in the form of a lecture. In this presentation, the results of their project should be presented and embedded in existing literature. In addition, students prepare a protocol in the form of a scientific paper.  
The practical course includes cellular and molecular aspects that are addressed in the model organism mouse. This includes the following work: Basic techniques of mouse genetics and experimental surgical methods, processing of brain tissue for immunohistochemistry, basics of working with primary cell cultures, immunofluorescence microscopy, confocal microscopy and biochemical techniques.  
Primary cell culture experiments are used to analyze techniques such as phagocytosis efficiency of different cell types. Immunohistochemistry is used to analyze cell-specific markers in the different disease states. Microscopy allows us to record the cellular and systemic events.
Students also have the opportunity to observe surgical methods such as experimental stroke surgery if they wish.

Educational objectives / Competences

Students master the basic techniques used in research into neurodegenerative diseases, among other things. The different methods allow us to ask specific questions. Accordingly, students name the advantages and disadvantages of different model systems. Students evaluate experimentally obtained data and use image processing and analysis software independently. Students write a scientific paper, present their results and communicate them in an international environment.

PD Dr. Florian Freudenberg

Contents

 In this practical course, the molecular and cellular causes of psychiatric disorders are investigated. Students encounter a wide range of translational methods. This includes cell culture techniques for the functional investigation of candidate genes (including the  production of primary cell cultures from humans and/or mice, production of viral vectors and viral gene transfer) and the investigation of molecular and cell biological mechanisms in the cell model and/or optionally in the mouse model (including the behavioral biological investigation of mice that have been genetically modified and/or pharmacologically treated). Following such experiments, various molecular biological (including quantitative PCR, Western blot, ELISA), neuroanatomical (brain dissection, cryostat sectioning, staining methods), immunohistochemical and microscopic (including fluorescence microscopy with and without structured illumination) characterizations will be performed. There is also the opportunity to gain insight into behavioral studies and imaging techniques (functional magnetic resonance imaging [fMRI], electroencephalography [EEG],
magnetoencephalography [MEG]) as used to assess abnormal neural processing in human psychiatric disorders.
The experiments are carried out in the laboratories of the Department of Psychiatry, Psychosomatics and Psychotherapy at Frankfurt University Hospital. Current projects of the clinic are worked on under supervision. The results are presented as part of a lecture in the laboratory seminar and documented in the form of a protocol.

Educational objectives / Competences

Students plan, conduct and analyze experiments that are used to investigate psychiatric disorders. Students develop scientific approaches and literature research. Students document their results and communicate them in oral and written form. In the seminar series (including the opportunity to participate in case presentations), students also acquire basic knowledge of the psychiatric disorders studied and are able to describe them.

PD Dr. Stefan Momma

Contents

Extracellular vesicles are membrane vesicles secreted by cells of all living organisms that can contain functional proteins and lipids as well as nucleic acids. Communication between cells through extracellular vesicles is a relatively new topic with high relevance in a variety of fields. In this practical course an introduction to the biology of extracellular vesicles is given.  The focus is on aspects of purification and classification, analysis of RNA/protein content as well as visualization and analysis of the transfer of functional molecules between cell populations in vitro and in vivo. Methods to be used in the practical course are e.g: Cell culture techniques, immunofluorescence microscopy, flow cytometry and other related techniques. The basics are formed through independent literature work and discussion of current publications. The results of the practical course are presented as a short lecture.

Educational objectives / Competences

The student gains an initial insight into this very young field of the biology of intercellular communication through extracellular vesicles. The student masters basic techniques for working with these vesicles and analyzing their biological functions. The experiments are carried out in coordination with current scientific studies, so that the student will be able to plan and carry out more complex scientific questions.

Prof. Dr. Andreas  Geburtig-Chiocchetti

Contents

This module teaches the basics for deciphering the molecular genetic mechanisms behind neuropsychiatric disorders such as autism or social behavior disorders. On the one hand, bioinformatic methods can be learned to identify genetic markers for the disorders or to link transcriptome data from neuronal cell models with patient data so that new findings about the cause of the disorder or biologically defined subgroups of patients can be identified. In particular, omics analyses and machine learning methods are used here. On the other hand, neuronal human cell models are generated based on the identified genetic markers (CRISPTR/Cas9). Students learn how to work with human cell models, in particular human embryonic stem cells (hESCs), induced pluripotent stem cells and especially the cultivation and quantitative analysis (real-time PCR, confocal microscopy) of neuronal progenitor cells, differentiated neurons and cerebral organoids. The participants will discuss the results during the working group's scientific seminar.

Educational objectives / Competences

The students implement translational research approaches. In particular, they use molecular genetics and bioinformatic methods to link findings from the cell model with patient data and vice versa.

Prof. Dr. Leo Kurian

Contents

This module provides a comprehensive exploration of the biology of human pluripotent stem cells (hPSCs) and their applications in programming cellular identity, with a particular focus on cardiovascular development and disease  modeling.

Theory:
This module covers the basic concepts of human pluripotent stem cell biology, focusing on how cellular identity is programmed and reprogrammed. Students will explore nuclear reprogramming mechanisms, along with direct and indirect methods of cell fate conversion. A significant portion of the module is dedicated to programming cardiovascular cell fate and applying human induced pluripotent stem cells (hiPSCs) for disease modeling. The module also delves into advanced topics such as the creation of organoids and ex vivo embryo models.
Practical Work:
Students will gain hands-on experience in culturing and handling hiPSCs, as well as differentiating them into cardiomyocytes in 2D culture systems. They will also learn how to generate cardiac organoids and apply functional genomics of RNA regulatory elements using hiPSC-based models. This includes the use of CRISPR/Cas9 based loss of unction technology to study gene function, targeting RNA-binding proteins and non-coding RNAs, and introducing mutations or endogenous tags. Additional practical work includes the creation of reporter cell lines, design and execution of arrayed screens, and investigating the role of context-specific splicing and selective translation mechanisms in regulating embryonic cell fate and cell identity.

Educational objectives / Competences

Students have a solid foundation on the basics of human pluripotent stem cell biology. Further. they have an advanced knowledge of modeling early embryonic development, cardiac development, genome engineering, and RNA regulatory principles of embryonic fate decisions.

Dr. Phillip Grote

Contents

This module serves as an introduction to the function of non-coding RNAs in the physiological context with a focus on tumor biology. The main model used in our lab is the mouse model system, which is preceded by detailed work in cell culture in vitro. The student will carry out a sub-project of current research in the lab. The focus of the lab is on the involvement of long, non-protein-coding RNAs (lncRNAs) in various cell regulatory processes in the bone marrow tumor microenvironment. For this purpose, CRISPR/Cas9 is used to genetically manipulate tumor cells as well as tumor-associated cells of humans or mice to generate genetic mutants of lncRNAs. In addition, we use other CRISPR technologies (e.g. CRISPRa (gene activation) and CRISPRi (gene inactivation)) to modulate gene programs in vitro. Such genetically modified cells are then tested in tumor microenvironment simulating co-cultures and can later be used in mouse systems.
At the end of the practical course, students will prepare a short written report (protocol) and present their findings to their lab colleagues in the lab seminar for discussion and possible future research direction. Students will participate in the weekly lab seminars and familiarize themselves with current research in the lab.

Educational objectives / Competences

Students use various CRISPR systems to study gene regulatory programs in tumor cells and tumor-associated cells and apply this to any other area of research. They "read" and interpret whole genome data to formulate hypotheses about gene regulation and design genetic manipulation experiments to verify the hypotheses.

Prof. Dr. Maik Böhmer

Contents

This laboratory course provides students with hands-on experience in advanced microscopy techniques and computational methods for extracting quantitative insights from complex biological imaging data. The first half of the course focuses on high-content screening (HCS) using automated multiwell confocal microscopy. Students will learn to transform microscopy images into quantitative data using machine learning and deep learning-powered image analysis tools. As a case study, reporter analyses in plant systems will be examined, specifically focusing on calcium measurements in Arabidopsis roots. Students will have the opportunity to conduct time-series analyses, investigate mutant studies, and perform pharmacological screens while gaining experience with a high-throughput spinning disk microscope. In the second half of the course, Arabidopsis root systems will be analyzed under physiological conditions using advanced light sheet microscopy. Students will track the movement of root amyloplasts during gravitropic responses and have the opportunity to design and 3D-print custom hardware to support these experiments. Throughout the course, they will work on cutting-edge research projects under the supervision of experienced postdoctoral researchers. The course will finish with the preparation of a comprehensive protocol and a seminar presentation to share their findings with the laboratory members.  

Educational objectives / Competences

Students know the basic concepts of quantitative image analysis using high-content screening confocal and light sheet microscopy. They apply computational image analysis techniques, including machine learning, deep learning, quantitative data analysis, and visualization for biological applications. Furthermore, students will gain experience with and apply optogenetic tools, and reporter assays in plant systems. They develop practical skills in experimental 
design, data collection, and scientific communication.