# data[].attributes.body
• How can I use this?

Flow cytometry allows for very fast accumulation of quantitative data in cellular analysis. In the past I have used flow cytometry for immune profiling of mouse and human blood samples, as well as cell cycle experiments. Flow cytometry is an essential tool in immunology research as well as in many other fields. For example, in the field of stem cell or tissue regeneration research, flow cytometry can be used to quickly determine the surface marker expression profiles of cells, which in turn can be used to determine things such as differentiation efficiency.

• Why must I use this?

There are times when large scale quantitative data must be presented in order to provide more compelling evidence for trends or status of cell populations. In times when other techniques, such as imaging methods, can only provide a narrow view of the situation, flow cytometry is required to provide a macro view of the phenotype of cells.

• When will I use this?

Flow cytometry is used when requiring relatively fast and high volume acquisition of data on cell populations. When I want to see the expression profiles of a group of cells and how this changes during my experiments.

# data[].attributes.headline
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.body
Learning this lecture changed how I view C. elegans. I can actually use this model—not just describe it. The worm gives me rapid and cheap genetics: I can create mutants, apply RNAi by feeding, and generate transgenic lines much faster than in mice. This matters because I can test my ideas within 2–3 weeks instead of months.

I use C. elegans when the question is about conserved cell biology or signaling pathways. The apoptosis pathway discovered in worms and later confirmed in humans is a real example of why data from worms can translate. Also, the transparent body and invariant cell lineage mean that if I want to track cell division or neuron behavior, worms are more informative and realistic than cultured cells.

I will use this model especially at early phases of a project: screening candidate genes, checking phenotypes, and doing rapid loss-of-function (RNAi/CRISPR) before committing to expensive mammalian work. In short, C. elegans is a practical decision—cheap, fast, genetic-powerful—and helps me fail early or succeed early.

# data[].attributes.headline
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.body
From this lecture, I realized that I should combine RNA techniques based on purpose. I will use RNA-Seq to discover candidate genes, qPCR to validate expression changes, and Northern blot when I must confirm real transcript size or splicing forms. RNA-FISH will be useful whenever localization matters, and RIP/EMSA will help prove RNA–protein interactions.

I use these methods because each reveals a different layer: quantity (qPCR), authenticity (Northern), spatial distribution (FISH), and mechanism (RIP/EMSA).

I will apply this toolkit when designing future experiments: first, screening with high-throughput data, then confirming expression with qPCR, and finally moving to structural and mechanistic assays only when necessary. This strategy saves time, reduces cost, and produces solid, believable data.

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1. How can I use this?

I can use C. elegans as a simple living model to study how genes and metabolic signals affect a whole organism. With tools such as gene knockdown, transgenic expression and fluorescent imaging, I can observe how cells develop, communicate and respond to changes in metabolism. These methods allow me to see biological effects at the organism level, rather than only in isolated cells.

2. Why must I use this?

I should use C. elegans because it offers a clear and practical way to study complex biology in a living system. It is easy to grow, genetically well understood and transparent, which makes it possible to directly observe biological processes in real time. Even though it is simpler than mammals, it still captures many conserved metabolic and signaling pathways. This makes it useful for gaining basic insights that can later inform more complex studies, including projects related to metabolic regulation.

3. When will I use this?

I will use C. elegans when I need in vivo information but want a faster and more accessible model than larger animals. It is especially useful during early-stage experiments, when I am exploring how genetic or metabolic changes affect overall physiology. If I need to test general effects of metabolites or gene regulation in a whole organism before moving to advanced models, C. elegans provides an efficient starting point.

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• How can I use this?

I can use mouse models for cancer research. In the past my lab had used a mutant mouse prone to develop diabetes, along with another mutant prone to develop pancreatic cancer, to conduct research into the relationship between diabetes, chronic high blood glucose, and the development of pancreatic cancer. The diabetes prone strain of mouse was an established mutant strain with a mutation in it's leptin receptor, named db mice. The pancreatic cancer prone strain of mouse used transgenic methods to produce a mouse line carrying the Kras G12D mutation. The two strains of mice were then crossed to produce mice that were both diabetic and carrying a mutation making them prone to develop pancreatic cancer. In this way, experiments for obtaining insights into the relationship between diabetes and pancreatic cancer could be carried out. Transgenic mice can also have "reporter genes" engineered into their genome, so that colors or patterns in their fur coat can be used to discern if they are indeed carrying the gene of interest, in the event they are crossed with another strain of mouse. This can partially save time in regards to genotyping the mouse.

• Why must I use this?

In many areas of research, especially in topics related to a clinical setting, experiments need to be conducted on systemic models rather than on individual component units such as cells. For example in the study of cancer a researcher may be interested in finding out the time frame in which metastasis occur. This type of observation is hard to reproduce accurately without animal models. When mere cell culture experiments are insufficient, animal models are required, and mice are both similar enough to humans, while being easy to maintain to make them one the most suitable model organisms.

• When will I use this?

I will use mouse models, or any other animal models, when mere cellular level experiments are insufficient for the research topic at hand. If my research is trying to demonstrate biological phenomena whose data will later to be extropolated to humans, and which can only be observed in a systemic setting, then mouse or animal models will be indispensable if I want my experiments to be convincing.

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How can I use C. elegans?

I will use C. elegans as an in-vivo model to validate and extend findings obtained from in-vitro cell-based assays. Specifically, it will help me evaluate organism-level effects such as survival, stress resistance, oxidative stress response, immune modulation, and lifespan after treatment with bioactive compounds or probiotic-derived factors. Using C. elegans allows me to connect molecular and cellular mechanisms (e.g., antioxidant or anti-inflammatory effects) with physiological outcomes in a whole organism.

Why I Must Use C. elegans?

While cell culture models provide mechanistic insights, they cannot fully represent systemic biological responses. C. elegans is essential because it offers a simple yet genetically conserved organism in which key pathways related to inflammation, oxidative stress, metabolism, and aging are shared with humans. It is cost-effective, ethically favorable, and enables rapid experimentation, making it an ideal intermediate model before moving to complex mammalian systems. Using C. elegans therefore strengthens the biological relevance, credibility, and translational value of my research.

When I Will Use C. elegans?

I will use C. elegans:

After obtaining significant results from in-vitro assays (e.g., antioxidant, cytotoxicity, or anti-inflammatory studies)

During mechanistic studies to evaluate whole-organism responses such as stress tolerance and lifespan

In preclinical validation of probiotics or bioactive compounds

While preparing publications or research proposals that require in-vivo evidence to support cellular findings

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• How can I use this?

By growing tissue organoids, 3D self-organizing structures derived from stem cells, I can model complex tissue structures and functions, allowing me to simulate cancer progression, tumor microenvironments, or developmental processes in vitro for drug testing and bio-pathway studies.

• Why must I use this?

Traditional 2D cell cultures fail to capture spatial organization, cell-cell interactions, and heterogeneity seen in vivo, while organoids provide physiologically relevant models that bridge the gap between simplistic monolayers and animal studies.

• When will I use this?

I will use organoids when investigating cancer metastasis, screening therapies for patient-specific tumors, or studying embryonic development, especially to predict drug responses or dissect genetic regulatory networks in a human-relevant context.

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• How can I use this?

Where I work, biotech or pharmaceutical companies often use high-throughput screening with automated imaging systems to process multi-well plates of cell cultures, in order to test drug or compound effects on cellular responses, such as viability, morphology, or signaling, enabling rapid evaluation of thousands of conditions simultaneously.

• Why must I use this?

Manual screening is too slow and labor-intensive for large compound libraries, while automated imaging provides quantitative, reproducible data on diverse phenotypes, uncovering subtle drug responses that low-throughput methods might miss. Automated systems also allows for the removal of human error as a factor.

• When will I use this?

I will use these methods when screening chemical libraries for hits in drug discovery, validating compound toxicity or efficacy across cell types, or assessing genetic perturbations' effects on cellular behavior in a scalable and reproducible way.

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How can I use this?

I can use C. elegans to study genetics, development, aging, and neural circuits because it’s transparent, easy to manipulate, and has powerful genetic tools like RNAi and CRISPR.

Why must I use this?

Because C. elegans gives you fast, cheap, and reliable insights into biology that often translate to humans—its genome shares 50–80% homology, and many pathways (like apoptosis) are conserved.

When will I use this?

I’ll use it when I need a simple but powerful system to test gene function, model human diseases, or run large-scale genetic screens—especially in projects where speed and cost matter.

# data[].attributes.headline
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

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I use genetically modified mice when I need clear, causal answers that cell culture can’t give me. Techniques like CRISPR knockouts, knock-ins, or conditional models let me test what a gene actually does in a real physiological setting, where immune cells, tissues, and metabolism all interact. I use these models when I want something close to human disease, when in vitro results aren’t convincing, or when I need to track cells or gene expression in vivo. Overall, they give me the most reliable and publishable way to connect a gene to a real biological function.

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• How can I use this?

These strategies allow me to simultaneously test multiple gene perturbations or interactions within gene circuits by using high-throughput genetic manipulation tools like CRISPR libraries. This helps identify key genes or pathways involved in complex biological processes or phenotypes.

• Why must I use this?

Traditional one-gene-at-a-time screens are time-consuming and miss combinatorial effects. Multiplexed screens enable efficient discovery of gene interactions and synthetic genetic relationships, which is critical for understanding gene networks and designing engineered gene circuits.

• When will I use this?

I will use these methods when studying complex traits influenced by multiple genes, mapping interaction networks, or optimizing synthetic biology constructs that require coordinated regulation of several genes simultaneously.

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• How can I use this?

DNA biosensing can be used in many different ways in bioscience research. From the earliest and simplest use of probes such as FISH, to modern multi-parameter large scale single cell or tissue genomics studies, DNA biosensing allows either focused or wide ranging studies with high throughput.

• Why must I use this?

Sometimes genome wide observations must be made in order to understand the whole picture in regards to the genomic profile either at the tissue level or at the cellular level. DNA probes or DNA sensing techniques allow this to be done in a high precise and specific manner.

• When will I use this?

When I want to see multi-parameter data using DNA probes. For example, in my lab, there has been projects in the past that try to find the most ideal conditions for differentiating iPSCs into specific tissue organoids. By designing a DNA aptamer biosensor to measure the growth factor release, I can also learn more about the signaling patterns of differentiated cells during organoid development.

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• How can I use this?

RNA techniques, such as RT-PCR and RNA sequencing, allow me to detect and quantify gene expression in cells. These methods can be applied to study how genes respond to different treatments or during development by analyzing mRNA levels, providing insight into cellular functions and regulatory pathways.

• Why must I use this?

Understanding gene expression helps reveal which genes are active under specific conditions. RNA techniques provide sensitive and precise ways to measure the RNA transcripts, which is key to linking gene activity with specific cellular responses, or disease states.

• When will I use this?

I will use RNA techniques when I want to analyze gene expression changes, validate gene knockdown or overexpression, or profile transcriptomes to discover new RNA sequences or investigate alternative splicing patterns in various biological contexts.

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Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

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How can I use this?
I can use C. elegans as a simple multicellular model to study how Bifidobacteria or their metabolites affect host metabolism and GLP-1 signaling pathways. By feeding worms with specific bacterial strains or metabolites, I can monitor changes in gut function, metabolic gene expression, and longevity, using fluorescent reporters or behavioral assays.

Why must I use this?
C. elegans provides a fast, cost-effective, and genetically tractable system to study host–microbe interactions in vivo. Its conserved metabolic pathways allow me to model aspects of human GLP-1 regulation and predict the systemic effects of probiotic interventions before moving to more complex models.

When will I use this?
I will use C. elegans in the early-to-mid stages of my project, after in vitro screening of bacterial strains. This system will help identify promising probiotic candidates and their mechanisms of action, guiding subsequent studies in organoids or mouse models.

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Nguyen Van Tat Thanh 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh 313302026 - tonion.ls13@nycu.edu.tw

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How can I use this?
I can use transgenesis and gene targeting to create mouse models that carry specific Bifidobacteria-responsive genes or GLP-1 reporters. Techniques like CRISPR/Cas9 or traditional homologous recombination allow me to insert, delete, or modify genes in mice to study how bacterial metabolites influence GLP-1 expression in vivo.

Why must I use this?
Mouse models are essential for validating findings from in vitro or organoid experiments. They allow me to observe systemic effects of probiotics on metabolism, GLP-1 regulation, and glucose homeostasis in a whole-organism context, which cannot be fully captured in cells or organoids.

When will I use this?
I will use transgenic or gene-targeted mice after identifying key Bifidobacteria strains and their target pathways in vitro and in organoids. These models will help confirm mechanisms and evaluate the therapeutic potential of probiotics in regulating GLP-1 and reducing diabetes-related phenotypes.

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Nguyen Van Tat Thanh - 313302026- tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026- tonion.ls13@nycu.edu.tw

# data[].attributes.body
How can I use this?
I can use organoid technology to test how Bifidobacteria strains influence GLP-1 expression and host metabolic responses in a controlled 3D environment. By co-culturing probiotics or their metabolites with intestinal organoids, I can measure changes in GLP-1 secretion, epithelial cell signaling, or metabolic gene expression, mimicking the human gut more accurately than 2D cultures.

Why must I use this?
Organoids retain the architecture, heterogeneity, and functional properties of human intestinal tissue, which are essential to study complex host–microbe interactions. They allow testing probiotic effects in a physiologically relevant setting before moving to animal models, reducing uncertainty and improving translational relevance.

When will I use this?
I will use organoid co-cultures in the mid-stage of my project, after identifying promising Bifidobacteria strains from initial screenings. This platform will help validate their impact on GLP-1 regulation and host metabolism before testing in vivo.

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Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.body
How can I use this?
I can use high-throughput screening to systematically identify Bifidobacteria genes or secreted factors that influence GLP-1 expression. By combining genetic libraries or metabolite libraries with cell-based assays that measure GLP-1 secretion or activity, I can rapidly test many candidates in parallel and detect those with the strongest effects.

Why must I use this?
GLP-1 regulation is controlled by complex interactions among bacterial genes and metabolites. Low-throughput approaches would be too slow to capture all potential contributors. High-throughput screening provides a scalable, unbiased way to discover key regulators and prioritize candidates for further study.

When will I use this?
I will use HTS during the early discovery phase to identify bacterial genes or compounds that modulate GLP-1. Hits from these screens can then be validated in functional assays and used to design engineered probiotics with enhanced metabolic benefits.

# data[].attributes.headline
313302026 Nguyen Van Tat Thanh - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
313302026 Nguyen Van Tat Thanh - tonion.ls13@nycu.edu.tw

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How can I use this?
I can use protein structural biology to study how the 3D structure of Bifidobacteria proteins relates to their role in regulating GLP-1 expression. Techniques like X-ray crystallography, NMR, or cryo-EM allow me to see how specific amino acids affect enzyme activity or interactions with host molecules. Structural insights can guide the design or modification of proteins to enhance their stability or function in engineered probiotics.

Why must I use this?
Protein function often depends on correct folding and interactions. For my project, understanding the structure of key bacterial proteins is crucial to predict how they influence GLP-1 signaling and metabolic regulation. Structural knowledge also helps identify residues for targeted modifications to improve probiotic efficacy.

When will I use this?
I will use structural biology after purifying proteins from selected Bifidobacteria strains. Comparing natural and engineered proteins will show how structural changes affect activity, helping optimize probiotic strains for controlling GLP-1 expression.

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Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

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How can I use this?

I can apply combinatorial and multiplexed genetic screening to systematically investigate which genetic features of Bifidobacteria drive GLP-1 induction in host cells. Instead of modifying one gene at a time, I can create a pooled library of Bifidobacterium variants—each carrying different combinations of metabolic genes, surface proteins, or regulatory elements suspected to interact with host enteroendocrine pathways.
By introducing DNA barcodes into each variant and co-culturing the entire library with intestinal organoids or GLP-1–reporter cell lines, I can track which genetic combinations most strongly activate GLP-1 production. This allows me to map synergistic or antagonistic gene interactions that shape how Bifidobacteria influence incretin signaling.

Why must I use this?

GLP-1 regulation by gut bacteria is not controlled by a single factor. It depends on complex metabolic processes such as SCFA production, bile acid modification, carbohydrate utilization pathways, and bacterial surface molecules that interact with host receptors. These factors often act in combination rather than isolation.
Using single-gene approaches would be too slow and would miss synergistic effects. Multiplex genetic screening gives me a high-throughput and unbiased strategy to pinpoint which combinations of bacterial genes—rather than individual ones—are necessary and sufficient to stimulate GLP-1.
This is essential for my research goal, because identifying the right gene networks will help engineer or select probiotic strains with maximal antidiabetic function.

When will I use this?

I will use this approach during the discovery and optimization phase of my project, when I need to identify which bacterial pathways contribute most to GLP-1 induction. Initially, I can perform in vitro screening using GLP-1 reporter intestinal organoids to rapidly narrow down candidate gene combinations.
After selecting the top-performing variants from the pooled screen, I will test them individually in vivo in diabetic mouse models to validate their effect on GLP-1 levels and glucose metabolism.
In the long term, combinatorial screening will be helpful if I work on engineering next-generation Bifidobacteria probiotics or uncovering compensatory pathways that determine which strains are most effective for diabetes prevention.

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Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.body
How can I use this?

I can use DNA nanotechnology to design highly programmable biosensors based on predictable base-pairing and precise structural assembly. DNA can be engineered into 2D or 3D nanostructures—such as DNA origami, tiles, or nanomachines—that respond to specific biomolecules. For example, DNA aptamers can be integrated into these nanostructures to recognize glucose, inflammatory markers, or microbial metabolites. When a target binds, the DNA structure changes conformation or produces a measurable signal, enabling sensitive detection.

Why can I use this?

I can use DNA nanotechnology because DNA is remarkably stable, easy to synthesize, and capable of self-assembly with high precision. These properties allow me to build customizable sensing platforms that are cheaper and more adaptable than antibody-based systems. DNA sensors also support multiplex detection, rapid signal generation, and integration with optical or electrochemical readouts, making them suitable for biomedical diagnostics and point-of-care applications.

When will I use this?

I will use this approach when I need to detect low-abundance biomarkers or design highly specific recognition systems. In metabolic or microbiome-related research, I could apply DNA aptamer-based sensors to monitor bacterial metabolites, inflammatory cytokines, or hormones such as GLP-1. This is especially useful for studying host–microbe interactions or developing diagnostic tools for metabolic diseases.

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Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.body
How can I use this?

I can use RNA-based techniques—such as qRT-PCR, RNA-seq, and small RNA profiling—to measure how Bifidobacteria or their metabolites influence GLP-1 expression in host intestinal cells. qRT-PCR allows me to quantify changes in GLP-1 mRNA after treating cells with bacterial supernatants, while RNA-seq helps identify broader transcriptional changes in pathways related to metabolism, inflammation, or enteroendocrine cell function. I can also analyze bacterial RNA to understand which genes are active when Bifidobacteria interact with the host.

Why must I use this?

RNA techniques are essential because gene expression changes are the most direct indicators of how Bifidobacteria regulate GLP-1. Without measuring RNA, I cannot determine whether bacteria enhance transcription, affect mRNA stability, or modify signaling pathways involved in diabetes control. RNA-seq is especially important for discovering new bacterial factors or host genes that mediate GLP-1 induction, providing mechanistic insight into microbe–host metabolic communication.

When will I use this?

I will use RNA techniques whenever I need to evaluate how bacterial treatments affect host gene expression—for example, after co-culturing Bifidobacteria with intestinal cells, testing engineered strains, or validating pathway activation. These methods are also crucial during hypothesis generation (RNA-seq screens) and during targeted validation steps (qRT-PCR) in my thesis project.

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Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.body
How can I use this?

I can use gene transfer to introduce specific genes, reporters, or regulatory elements into bacterial or mammalian cells so I can study how they influence GLP-1 expression. For example, I could transfer plasmids into Bifidobacteria to express candidate enzymes or metabolites, or use viral vectors to deliver GLP-1–related reporter constructs into intestinal cell models. This allows me to test how bacterial genes or secreted factors affect host signaling pathways involved in glucose regulation.

Why must I use this?

To understand how Bifidobacteria regulate GLP-1, I need a precise way to modify genes in both the bacteria and the host cells they interact with. Gene transfer is essential for validating whether a bacterial gene contributes to GLP-1 stimulation, for creating knockout or overexpression strains, and for testing regulatory elements in host intestinal cells. Without gene transfer, it would be impossible to dissect mechanisms or confirm causality in microbe–host metabolic interactions.

When will I use this?

I will use gene transfer whenever I need to alter gene expression in my experiments—for example, when engineering Bifidobacteria strains, testing mutants, creating GLP-1 reporter cell lines, or introducing CRISPR components to study pathway regulation. It will be especially useful during functional validation and mechanistic studies directly related to my thesis.

# data[].attributes.headline
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.body
How can I use this?

I can use protein structural biology to understand how a protein’s three-dimensional shape determines its function, binding specificity, and stability. By analyzing structures obtained from X-ray crystallography, cryo-EM, or computational models, I can identify key residues important for enzyme activity or host–microbe interactions relevant to my research. This knowledge helps me interpret how a protein behaves, predict its role in biological pathways, and detect structural problems such as misfolding in recombinant protein expression.

Why must I use this?

Protein structure is essential for explaining function at the molecular level. Small structural changes can completely alter activity, binding, or regulatory mechanisms. For my thesis, structural information is necessary to understand how bacterial or host proteins interact during infection or immune signaling. It also allows me to identify residues for mutagenesis, evaluate the effects of pathogenic variants, and design more stable or more active protein variants for experimental studies.

When will I use this?

I will use structural biology whenever I need to infer a protein’s function, evaluate binding interactions, or troubleshoot recombinant protein experiments. It becomes especially important after purifying a target protein, when I want to compare wild-type and mutant forms or understand how structural changes influence biological outcomes related to host–microbe interactions.

# data[].attributes.headline
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

# data[].attributes.subject
Nguyen Van Tat Thanh - 313302026 - tonion.ls13@nycu.edu.tw

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How can I use this?

I can use recombinant protein design to produce specific bacterial or host proteins in a clean, controlled form. By cloning a gene of interest into an expression vector and purifying the protein with tags like His-tag, I obtain the exact molecule I need for functional or interaction studies related to my research.

Why must I use this?

I need this technique because natural proteins are often difficult to isolate in large, pure amounts. Recombinant expression allows me to study how individual microbial or immune-related proteins function, interact, or contribute to processes like inflammation and sepsis—key topics in my thesis.

When will I use this?

I will use recombinant protein design whenever I need to test a protein’s activity outside the whole organism, such as examining how a bacterial factor binds to a host receptor or affects immune signaling. It is especially useful for biochemical assays and validation experiments in my Master’s research.

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How Can I Use This?

I can use the nematode Caenorhabditis elegans for multicellular research by employing methods such as microinjection and biolistic transformation to introduce foreign DNA. Microinjection is a common and efficient method where DNA is injected into the syncytial gonad to produce transgenic lines, often using a visible marker as a co-injection marker to identify transgenic worms. A major limitation of this technique is that the expression of the transgene can sometimes be silenced in germ cells. Alternatively, biolistic transformation involves shooting DNA coated gold particles into the worms, which generally results in stable, chromosomally integrated transgenes that do not undergo germline silencing, though this method is more labor-intensive.

Why Must I Use This?

I must use C. elegans to study biological principles due to its combination of simplicity, genetic tractability, and well-characterized development. It is highly favored over other multicellular models because it is transparent, allowing for easy visualization of all its internal structures, and has a fixed and fully mapped cell lineage from egg to adult. This complete cellular map is unavailable in any other animal. Furthermore, C. elegans is a hermaphrodite, which greatly simplifies genetic crossing and mutant isolation. Compared to more complex models like mice, C. elegans offers a fast and cheap system for large-scale genetic screenings.

When Will I Use This?

I will use C. elegans when your research question can be addressed with a model that is evolutionarily distant from humans but still shares fundamental biological processes. This is especially true when you need to study developmental biology, aging, neurobiology, or programmed cell death, as the core mechanisms of these processes are conserved. I will select this model if you need a fast lifecycle and high reproduction rate for efficient genetic screening or if you need a cost-effective alternative to mammalian models.

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How Can I Use This?

You can use these techniques through two principal methods: classical transgenesis and precise gene targeting. In transgenesis, the major method is pronuclear injection, where a DNA construct is manually injected into the male pronucleus of a one-cell embryo. The process involves collecting embryos, injecting the transgene, and implanting the embryos into a surrogate mother to guarantee the gene's presence in all cells of the resulting founder animal. Alternatively, gene targeting in Embryonic Stem Cells is used to create specific modifications like a knock-out or knock-in via homologous recombination. This involves modifying ESCs in vitro, selecting for the correctly targeted cells, injecting them into a host blastocyst to produce a chimeric mouse, and screening for germ-line transmission.

Why Must I Use This?

You must use genetically manipulated mice to accurately model the complex dynamics of human physiology and disease, which cannot be fully replicated in simpler organisms or cell cultures. Mice are favored because they share approximately 95% of their genes with humans, and their body systems function similarly, allowing researchers to study gene function, establish causation, and test therapies in a relevant mammalian context.

When Will I Use This?

You will use these methods whenever your research requires the comprehensive physiology of a whole mammal to investigate a gene’s function or model a complex human condition like diabetes or Alzheimer’s. You will use gene targeting when precision is critical, for instance, when creating an exact knock-in of a human disease-causing mutation. This system allows you to control the gene's deletion or expression spatially or temporally , thereby revealing its function in adult animals.

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How Can I Use This?

You can use this modeling technique by generating microscopic, self-organizing, three-dimensional organoids from stem cells. The foundational method involves taking either adult stem cells or pluripotent stem cells and culturing them to guide their self-assembly into structures that biologically and structurally resemble miniature human organs. Once these systems are established, the scale-up process is usually possible to allow for large-scale genomic screening or drug screening. Furthermore, you can apply most modern genetic engineering tools directly to these stem cells or organoid systems to study gene function or disease mechanisms.

Why Must I Use This?

You must use this modeling approach to create a better representation of human biology in a dish, as organoids offer the best recapitulation of human physiology compared to simple two-dimensional cell culture, animal models like C. elegans or fish, and PDX models. They are highly favored because they enable personalization, meaning they can be grown from an individual's stem cells to test tailored therapies or study patient-specific disease progression. Additionally, while they are more costly than traditional cell lines, they provide a valuable, physiologically complex alternative that is generally less expensive than using mouse or others models for screening.

When Will I Use This?

We should use next-generation modeling when your research requires a complex, multi-cellular structure that more accurately mimics the natural conditions of a human organ. This method is ideal when you need to perform large-scale genomic or drug screening where a high-throughput format is necessary, but the simplicity of 2D cell lines would lead to irrelevant or misleading results. Furthermore, organoids are chosen when the primary objective is personalized medicine, using models derived from an individual patient to precisely study disease and test drug efficacy for that specific person.

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How Can I Use HTS?

You can use High-Throughput Screening (HTS) to quickly test many thousands of different substances, like small chemical compounds or genetic materials, to see if they affect a specific biological target, such as a protein or enzyme. The core methods rely on optical assays, meaning the result is measured by light, often using fluorescence or luminescence techniques because they are sensitive and easy to automate. These tests are done in tiny wells of specialized plates. The main steps involve: developing a test that works well on a small scale, using robotic systems to automatically move the target, test compounds, and reagents into the plates, reading the result in every well using a specialized plate reader and finally analyzing the massive amount of data to find the active compounds, known as "hits."

Why Must I Use HTS?

You must use HTS primarily to find new starting points for drug development or to identify crucial biological mechanisms that control cell function. It is necessary when the task involves analyzing hundreds of thousands or even millions of samples contained in modern chemical libraries. HTS is highly favored over traditional, manual lab methods because it offers enormous advantages in speed, scale, and cost-effectiveness. Traditional techniques could only test a few compounds per week, while HTS uses sophisticated robotics and automation to test tens of thousands of compounds per day. This speed drastically accelerates the discovery process, minimizes human error, and saves money by using much smaller volumes of expensive reagents.

When Will I Use HTS?

You will typically use HTS at the very beginning of a research or drug discovery project, specifically when you need to explore a very large number of potential solutions or effectors. The necessary condition for using HTS is having a well-defined target and a large, diverse library of compounds or genetic materials to screen against it. Essentially, HTS is needed whenever the sheer volume of samples makes manual testing impractical, too slow, or too costly.

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How can I use it?
Flow cytometry is a technique that allows us to study the physical and chemical properties of cells in suspension, one cell at a time. I can use it to measure cell size, surface, and intracellular proteins, DNA content, or cell viability. By labeling cells with fluorescent antibodies or dyes, I can detect specific markers or molecules using a laser-based system. Manipulating this technique involves preparing and staining samples correctly, setting up gates during data analysis, and adjusting parameters like fluorescence intensity or scatter. I can also use cell sorting to isolate specific populations for further experiments, such as cloning, culturing, or genetic analysis.

Why must I use this?

Flow cytometry is essential because it provides rapid, quantitative, and multiparametric analysis of thousands or even millions of cells in a very short time. It allows me to study the immune system in great detail, identify rare cell types, and analyze functional properties such as cytokine production, proliferation, or apoptosis. Compared to traditional microscopy or ELISA, it offers higher precision, speed, and the ability to analyze multiple markers at once. Technology can also be combined with fluorescent barcoding or bead-based assays to study many samples or cytokines simultaneously, saving both time and reagents.

When will I use it?
I will use flow cytometry in research that involves immunology, cell biology, or biomedical studies. It is particularly useful for characterizing immune cell populations, watching responses to infection or vaccination, and studying diseases like cancer or autoimmune disorders. In clinical or translational research, flow cytometry helps diagnose immune deficiencies, leukemia, or lymphoma by identifying abnormal cell markers. I would also use it when I need to track cell proliferation, detect apoptosis, or measure protein phosphorylation. Overall, this technique is necessary whenever I need to analyze complex cellular systems quickly and accurately at the single-cell level.

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313303017 - Pham Nhat Phuong Trinh - pnhatptrinh@gmail.com

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313303017 - Pham Nhat Phuong Trinh - pnhatptrinh@gmail.com

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How can I use this?

I can use C. elegans to study developmental and physiological processes in a whole organism because its transparency, invariant cell lineage, and mapped nervous system allow direct observation of tissues and behavior. With genetic tools such as mutants, RNAi, CRISPR, and transgenics, I can perturb gene function and efficiently connect molecular mechanisms to organism-level phenotypes.

Why must I use this?

Because C. elegans combines high genetic conservation with experimental simplicity. Its short lifecycle, small cell number, and low maintenance cost make it ideal for rapid studies of development, stress responses, apoptosis, and aging. The ability to perform large-scale genetic screens and apply RNAi conveniently provides advantages that many other systems cannot offer.

When will I use this?

I will use C. elegans when I need a fast and tractable model to evaluate gene function, perform genetic screens, or analyze behavior linked to defined neural circuits. It is especially useful for aging research: in my previous Aging course, I saw how its short lifespan and clear aging markers make lifespan and healthspan assays highly efficient. This makes the worm a practical first model before moving to more complex systems.

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NGUYEN THI MINH HANG - 313302027

How can I use this?

I can use multicellular organism methods in C. elegans to study development, tissue interactions, and whole-body responses in a controlled and genetically tractable system. Because the worm is transparent and has a fully mapped cell lineage, I can easily visualize how cells divide, migrate, and differentiate during growth. By combining fluorescent reporters, CRISPR-based editing, RNAi, and tissue-specific promoters, I can observe how genetic changes affect the nervous system, muscles, immune pathways, metabolism, or stress responses across the entire organism. This gives me the ability to test hypotheses about multicellular coordination, organ function, and gene regulation in vivo, rather than relying only on isolated cell culture systems.

Why must I use this?

I must use C. elegans for multicellular studies because it provides a whole-organism context that simpler models cannot offer. Unlike cell lines, the worm allows me to observe interactions between tissues, developmental timing, and systemic physiological responses, all of which are essential for understanding how a gene or pathway functions within a complete biological network. The worm is also ethical, inexpensive, and genetically well-annotated, enabling rapid screening and mechanistic studies that would be too slow, costly, or complex in mammals. In addition, the high evolutionary conservation between C. elegans and humans means that discoveries made in worms often translate into insights about human development, aging, immunity, and disease.

When will I use this?

I will use these methods when my research requires whole-organism evidence, such as determining how a mutation affects multiple tissues, whether a signaling pathway influences development, or how different organs coordinate during stress or infection. This is especially important when cell culture models fail to capture systemic effects or long-range communication between tissues. C. elegans is ideal for the early and intermediate phases of my project, when I need to evaluate multicellular phenotypes, perform genetic screens, or monitor organism-wide outcomes before moving on to more complex vertebrate models. It helps me identify which genes, pathways, or conditions are worth pursuing in higher organisms.

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NGUYEN THI MINH HANG - 313302027

How can I use this?

I can use transgenesis and gene-targeting methods in mice to introduce precise genetic changes, add reporters to visualize gene activity, or control when and where a gene is expressed. Techniques such as CRISPR knock-in/knock-out, ES-cell targeting, pronuclear injection, and Cre/loxP allow me to test the function of specific regulatory elements, coding mutations, or gene networks directly in a whole organism. These approaches let me study gene function in a physiological setting that reflects real tissue complexity.

Why can I use this?

I can use these methods because mice are a genetically tractable model with strong biological similarity to humans. Their genome can be modified in a stable and heritable way, making them ideal for disease modeling and functional testing. Mouse lines also offer temporal and cell-type specificity through inducible or tissue-specific Cre drivers, which is essential when global knockouts or early misexpression would cause lethality or mask the true phenotype. This flexibility allows me to study genes under conditions that more accurately reflect human biology.

When can I use this?

I can use these approaches when my research questions require in vivo validation rather than just cell-culture results. This includes situations where I need to confirm whether a regulatory variant truly affects expression in a specific tissue, or when I want to activate or delete a gene in defined neuronal or developmental populations. These tools become necessary when the biological process I’m studying, such as neural circuitry, development, or disease progression, depends on interactions that only occur within an intact mammalian system.

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elise.gascher.bt14@nycu.tw.edu

How can I use this?

You can use C. elegans as a model organism to study many biological processes in a simple and efficient way. For example, you can observe development because the worm is transparent, or analyze gene function using tools like mutants, RNAi, or CRISPR. You can also use transgenics to see where a gene is expressed or what happens when you overexpress it. Basically, the worm lets you test ideas about genetics, cell biology, or behavior in a whole organism.

Why must I use this?

Because C. elegans is one of the easiest organisms to work with, while still sharing many conserved pathways with humans (about 50–80% of genes). It grows fast, is cheap to maintain, and has a fully mapped cell lineage and nervous system. It’s perfect when you want to understand fundamental mechanisms like development, cell death, aging, or behavior. And since it’s transparent and small, experiments are simple and reproducible. So, it’s a very practical and powerful model for research.

When will I use this?

You will use C. elegans when you need a fast, simple and reliable system to study gene function or biological pathways. For example, when you want to screen for mutants, test RNAi effects, analyze neuronal circuits, or explore aging in a short time. It’s also useful when you need large populations, or when working with mammals would be too slow, expensive, or ethically complex. Basically, anytime you need a genetic or developmental model that gives quick and clear results.

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elise.gascher.bt14@nycu.tw.edu

How can I use this?
You can use these techniques to change the mouse genome in a precise way. For example, you can add a transgene by pronuclear injection, or you can delete or modify a gene using gene targeting or CRISPR/Cas9. You can also control when and where a gene is active using systems like Cre/loxP or inducible promoters. Basically, these tools let you study gene function directly in a living organism.

Why must I use this?
Because mice are very similar to humans genetically, and we cannot do genetic experiments on humans for ethical reasons. Using genetically modified mice helps us understand how genes work, model human diseases, and test new therapies. It’s also one of the best ways to see the real effect of a gene in a whole organism, not just in cells.

When will I use this?
You will use these methods when you need to study the effect of a specific gene, create a disease model, or test how changing a gene influences development or physiology. For example, if a gene causes early lethality, you might use conditional or inducible systems to delete it only later or only in one tissue. In general, anytime you need to explore gene function in vivo, these techniques become essential.

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C. elegans, a microscopic, transparent worm, is a popular model for studying multicellular organisms due to its simple body plan, well-defined cell lineage, and rapid growth. It shares many conserved biochemical pathways with humans, is easily grown, and is genetically tractable. Because of these benefits, we can examine host-microbe interactions, immunological responses, toxicity, and the effects of medication in a whole-organism setting using C. elegans, without having to deal with the moral and economic challenges associated with using animal models, such as mice.

1. How can I use this?

During chemotherapy-like stress, I can easily assess if my next-generation probiotics can enhance survival, lessen toxicity, or alter host immune responses using C. elegans. For example, I can treat worms with my probiotic candidates after exposing them to chemotherapeutic treatments to see if they increase longevity, lower oxidative stress, preserve gut integrity, or alter immune-related signaling pathways. This provides me with an early indication of whether my germs have protective or damaging effects on the entire organism.

2. Why must I use this?

Compared to mammalian models, C. elegans allows me to look at host-microbe-drug interactions in a living multicellular organism much more quickly and ethically. It is affordable, genetically well-characterized, and scalable, providing me with biological complexity that cell culture cannot. Additionally, it enables me to effectively screen various probiotic strains and conditions for my study before proceeding with more expensive mouse experiments.

3. When will I use this?

C. elegans will be used in the early-stage screening phase, before moving on to mammalian cell tests and animal studies. It is ideal for determining which probiotic strains are worthwhile, evaluating toxicity, monitoring effects throughout the body, and choosing the most promising candidates to proceed to more complex in vitro or in vivo research.

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How, Why, and When to Use C. elegans as a Model Organism

How can this be used?

C. elegans enables controlled genetic and developmental studies because it is transparent, has an invariant cell lineage, and supports CRISPR editing, RNAi, and live imaging. Its short generation time and high fecundity make it suited for large-scale genetic screens, neuronal mapping, and studies of behavior, stress responses, and aging [1].

Why must this be used?

The worm shares extensive conservation with human disease pathways and allows mechanistic questions to be tested in an intact organism at single-cell resolution. Its simplicity, low cost, and compatibility with high-throughput phenotyping make it uniquely powerful for linking gene function to system-level outcomes [1,2].

When is this used?

C. elegans is selected for projects requiring rapid in vivo genetics, functional genomics, neurobiology studies, drug or toxicity testing, or modeling conserved signaling pathways before transitioning to vertebrate systems [2,3].

References

1.Calahorro F., Ruiz-Rubio M. Caenorhabditis elegans as an experimental tool for molecular neuroscience. Front Mol Neurosci, 2021.

2.Hunt P.R. The C. elegans model in toxicity testing. J Appl Toxicol, 2017.

3.Markaki M., Tavernarakis N. Modeling human diseases in Caenorhabditis elegans. Biotechnol J, 2020.

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Transgenesis and Gene Targeting in Mice, How, Why & When (Non-personal Version)

How can this be used?

Transgenesis and gene targeting in mice enable controlled manipulation of the genome to study gene function in vivo. Pronuclear injection introduces exogenous transgenes under defined promoters, while embryonic stem (ES) cell targeting and CRISPR-assisted homologous recombination generate precise knockouts, knock-ins, or reporter alleles, as outlined in the lecture slides. These methods create genetic models that reflect developmental, physiological, or disease-relevant contexts.

Why must this be used?

Mouse models reveal phenotypes and regulatory mechanisms that cannot be captured in cell culture. They allow analysis of tissue specificity, developmental timing, systemic interactions, and compensatory pathways critical factors for understanding gene function and modeling human disorders [1–3].

When will this be used?

These approaches are applied when validating pathogenic mutations, constructing cancer or immunology models, evaluating therapeutic strategies, or studying developmental processes that require an intact mammalian system before translational progression [2,3].

References

1.Capecchi M.R. Gene targeting in mice: functional analysis of the mammalian genome. Nat Rev Genet, 2005; 6: 507–512.

2.Yang H. et al. One-step generation of mice carrying reporter and conditional alleles by CRISPR/Cas-mediated genome engineering. Cell, 2013; 154: 1370–1379.

3.Platt R.J. et al. CRISPR–Cas9 knock-in mice for genome editing and cancer modeling. Cell, 2014; 159: 440–455.

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Next-Generation Modeling of Cancer and Development — How, Why & When

How can I use this?

I can use next-generation cancer and developmental models such as organoids and organ-on-chip systems to recreate human tissue architecture in vitro, allowing me to analyze tumor evolution, lineage specification, drug responses, and developmental signaling with far greater physiological relevance than 2D cultures. Using patient-derived organoids or iPSC-derived structures, I can model crypt–villus organization, airway differentiation, or brain development, These platforms can be combined with CRISPR editing, single-cell sequencing, and microfluidics to resolve dynamic behaviors at cellular and tissue scales.

Why must I use this?

These systems capture human-specific features that animal models often miss, including genetic diversity, self-organization, and clinically relevant drug sensitivity. Their scalability, compatibility with high-content imaging, and integration with omics make them essential for precision oncology and developmental research [1–3].

When will I use this?

I will use these models when validating patient-specific mutations, testing therapeutic candidates, mapping developmental defects, or studying host–pathogen interactions, especially where 2D or animal systems lack fidelity [2,3].

References

1.Clevers H. Modeling development and disease with organoids. Cell, 2016; 165: 1586–1597.

2.Sachs N. et al. A living biobank of breast cancer organoids captures disease heterogeneity. Cell, 2018; 172: 373–386. 3.KimJ., Koo B.-K., Knoblich J.A. Human organoids: model systems for human biology and medicine. Nat Rev Mol Cell Biol, 2020; 21: 571–584.

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High-Throughput Screening (HTS) — How, Why & When

How can I use this?

I can use high-throughput screening to test large chemical or genetic libraries in parallel using miniaturized assays, automated liquid handling, and plate readers. By building either target-based or phenotype-based assays, I can quantify enzymatic activity, signaling responses, viability, or morphological changes. A validated assay with a strong Z′-factor provides confidence that thousands of conditions can be screened reproducibly, allowing rapid identification of hits that modulate a pathway or phenotype [1,2].

Why must I use this?

HTS is essential when the biological question involves many potential regulators or compounds. It provides scalability, statistical robustness, and automation that manual experimentation cannot achieve. HTS also reduces bias by enabling unbiased discovery of unexpected targets, and it integrates seamlessly with downstream confirmation assays such as imaging or transcriptomics [2,3].

When will I use this?

I will use HTS in the early discovery phase screening chemicals, CRISPR libraries, or reporter systems to reveal initial hits. Later, I will apply it when comparing toxicities, optimizing dose–response behaviors, or prioritizing candidates for mechanistic studies [1,3].

References

1.Macarron R. et al. Impact of high-throughput screening in biomedical research. Nat Rev Drug Discov, 2011; 10: 188–195.

2.Arkin M.R., Whitty A. The road less traveled: high-throughput screening in academia. Nat Chem Biol, 2009; 5: 15–17.

3.Pushpakom S. et al. Drug repurposing: Progress, challenges and recommendations. Nat Rev Drug Discov, 2019; 18: 41–58.

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How can I use this?

Flow cytometry analyzes single cells suspended in fluid by measuring fluorescence as they pass through a laser beam. It quantifies surface or intracellular markers, while fluorescence-activated cell sorting (FACS) physically separates cells based on signal intensity [1,2]. This allows simultaneous measurement of multiple parameters at high speed and resolution.

Why must I use this?

It is essential for defining immune cell phenotypes, activation states, and cytokine production. Compared with bulk assays, flow cytometry provides single-cell precision and quantitative population distribution, while sorting yields pure subsets for downstream genomic or functional assays [2,3].

When will I use this?

I will use flow cytometry during immune profiling or infection studies, especially to analyze neutrophil or macrophage activation and to isolate specific subpopulations for transcriptomic or proteomic analysis [1,3].

References

1.Saeys Y., et al. Computational flow cytometry: Helping to make sense of high-dimensional immunology data. Nat Rev Immunol, 2016; 16: 449–462.

2.Herzenberg L.A., et al. The history and future of the fluorescence activated cell sorter and flow cytometry: A view from Stanford. Clin Chem, 2002; 48(10): 1819–1827.

3.Chattopadhyay P.K., Roederer M. A new era in cytometry: Multiparameter analysis of the immune system. Nat Rev Immunol, 2012; 12: 451–455.

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• How can I use this?

Methods in C. elegans can be used to study multicellular biology in a simple, easily manipulatable organism. I can use tools such as RNAi feeding, transgenic reporter strains, and cell-specific promoters to observe how genes regulate development, neural circuits, and behavioral patterns.

• Why must I use this?

Some biological processes like tissue development, cell-cell communication, and organism-level phenotypes, cannot be studied accurately in cell culture. C. elegans provides a transparent body, well-mapped cell lineage, and established genetic tools, making it ideal for linking gene function to physiological outcomes in a complete multicellular organism.

• When will I use this?

I will use these methods when I need an in vivo model to test how genetic changes affect development, behavior, or aging. For example, if I want to study how a mutation alters neuronal wiring, C. elegans methods allow me to observe these effects directly in a living organism.

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• How can I use this?

Gene transfer allows me to introduce specific DNA sequences into cells or organisms so I can express new genes, silence existing ones, or gene modifications. For example, I can use viral vectors or plasmids to deliver a reporter gene into mammalian cells to study signaling pathways or to test how certain regulatory elements affect gene expression.

• Why must I use this?

To study gene function, I need a way to precisely manipulate genetic material inside living cells. Many biological questions such as how a mutation alters phenotype, or how a therapeutic gene can correct a defect, cannot be addressed without introducing new genetic functions. Gene transfer provides an effective and efficient method to achieve this.

• When will I use this?

I will use gene transfer whenever an experiment requires changing what a cell expresses. This includes overexpressing proteins to study their roles, introducing CRISPR components for genome editing, creating stable cell lines that carry specific constructs, or delivering therapeutic genes in translational research.

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• How can I use this?

Protein structure biology provides the tools and concepts needed to understand how a protein's 3D conformation determines its function. In practical research, this allows me to examine how mutations alter protein folding, stability, and active-site geometry. For example, if I am studying an enzyme implicated in a disease pathway, determining its structure through X-ray crystallography, cryo-EM, or NMR spectroscopy enables me to map functional domains, locate catalytic residues, and visualize how ligands or inhibitors bind. With this information, I can rationally design experiments to test structure-function relationships, create targeted mutations, and screen for small molecules that may modulate the protein's activity.

• Why must I use this?

Protein behavior cannot be fully understood from its amino-acid sequence alone. Because many biological processes-signal transduction, metabolism, transcriptional regulation-depend on highly specific molecular interactions, knowing the structure is essential for explaining the mechanism at the atomic or near-atomic level. Structural information also provides insight into misfolding diseases, conformational changes, and protein-protein interfaces that are otherwise invisible using standard biochemical assays. Therefore, protein structure biology is necessary when the research question requires mechanistic clarity, such as determining how a mutation disrupts function, why an inhibitor works, or what conformational states exist during a protein’s activity cycle.

• When will I use this?

I will use protein structure biology whenever I need to visualize a protein's architecture to answer functional or mechanistic questions. For example, if I am working on drug development, structural data will guide rational design of molecules that fit precisely into a binding pocket. If I am studying a protein complex involved in immune signaling, cryo-EM can reveal how multiple subunits assemble and interact. Additionally, when mutations produce unexpected phenotypes, solving the structure can help explain how subtle changes reshape the protein’s stability or interaction surfaces. In short, I will use protein structure biology in any project where understanding the shape of the molecule is key to understanding its biology.

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1. How can I use this?
C. elegans is a transparent nematode with a short life cycle and a well-described anatomy. It can be genetically manipulated using mutagenesis, RNA interference, transgenesis, and CRISPR/Cas9. Researchers use it to study development, cell lineages, neuronal circuits, programmed cell death, and aging. Because of its simplicity and reproducibility, it is one of the most accessible multicellular models for laboratory studies.

2. Why must I use this?
C. elegans is a powerful model because it shares 50–80% of its genes with humans, yet it is easy and cheap to maintain. Its invariant cell lineage and compact genome make it ideal for genetic studies. Discoveries in C. elegans have revealed conserved pathways, such as apoptosis, that are directly relevant to human biology and disease. This means that results obtained in worms often provide insights that can be applied to higher organisms, including humans.

3. When will I use this?
This organism is used in projects on genetics, neurobiology, developmental biology, and aging. It is mobilized when researchers need a simple multicellular model with strong genetic tools, short generation time, and reproducible results. It is particularly useful when large-scale genetic screens or functional studies must be performed quickly and at low cost.

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1. How can I use this?
Transgenesis and gene targeting in mice allow scientists to introduce, delete, or modify genes in order to study their function. Methods include pronuclear injection, embryonic stem cell targeting, and somatic cell nuclear transfer. More recent approaches use CRISPR/Cas9 for precise gene editing. These techniques make it possible to create mouse models that carry transgenes, knockouts, or conditional alleles. In practice, this means researchers can design experiments that directly test how genes influence development, disease, or therapy response.

2. Why must I use this?
These methods are essential to understand gene function in living organisms, to model human diseases, and to test therapies before applying them to humans. They also allow the creation of animals with useful traits, such as producing therapeutic proteins or serving as organ donors. Without these approaches, it would be impossible to study gene expression in a controlled and reproducible way. Moreover, they provide a bridge between basic genetic research and translational applications in medicine.

3. When will I use this?
They are used in biomedical research whenever gene function must be studied in vivo. Applications include disease modeling (cancer, diabetes, cystic fibrosis, Alzheimer’s), functional genomics, and translational medicine. They are mobilized in genetics laboratories, molecular biology, and biotechnology. In addition, they are often chosen when researchers need reliable animal models that closely mimic human physiology.

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1. How can I use this?
High-throughput screening (HTS) methods allow testing thousands of compounds or genetic targets in parallel. They rely on automation, robotics, and sensitive detection systems. HTS can be used to identify active molecules, study gene functions, or discover new therapeutic candidates. By combining speed and scale, HTS makes it possible to explore complex biological questions in a systematic way.

2. Why must I use this?

HTS is indispensable for drug discovery, functional genomics, and biomarker identification. It provides large-scale data that help select promising compounds or genes for further study. Without HTS, biomedical research would be slower and less efficient. It also ensures that potential therapies can be evaluated early, reducing the risk and cost of later clinical trials.

3. When will I use this?

These methods are applied in pharmaceutical research, genetic studies, and biotechnology projects. They are mobilized whenever large numbers of samples or targets must be tested quickly and systematically. HTS is especially important when researchers need to compare many different molecules or genetic variants under standardized conditions.

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1. How can I use this?

Combinatorial and multiplexed genetic screens allow me to test many genes or genetic combinations at the same time, instead of one by one. By using technologies such as CRISPR libraries, barcoded genetic constructs or pooled screening approaches, I can efficiently explore which genes interact to influence a desired outcome. In my probiotic project, these strategies can help identify combinations of genes that most effectively modulate metabolite production or optimize metabolic pathways in engineered strains. This high-throughput approach saves time and resources while revealing interactions that would be difficult to detect otherwise.

2. Why must I use this?

I must use combinatorial and multiplexed screens because metabolic traits are often controlled by multiple genes working together. Testing individual genes separately may miss important synergies or conflicts between pathways. These strategies allow me to map out genetic interactions systematically and discover key regulators that drive metabolite changes. They also provide robust datasets that improve confidence in identifying the best genetic designs for my engineered probiotics which ensure that the final strains perform reliably under complex biological conditions.

3. When will I use this?

I will use these strategies during the early and middle stages of my project, when I need to explore a wide range of genetic possibilities. Early on, multiplexed screens can identify promising gene targets for metabolite modulation. Later, combinatorial approaches can help optimize interactions between multiple genes to maximize efficiency and stability of metabolite production. By using these strategies before moving to animal models, I can select the most promising probiotic designs and reduce experimental trial and error in downstream testing.

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In this week’s class, we learned about C. elegans as a model organism, including its anatomy, genetic tools, and why it is widely used in developmental and molecular biology. The transparency, short lifecycle, and strong genetic tool kit made me think about how worms could potentially complement my work on Acinetobacter baumannii antimicrobial resistance, especially when looking at host–pathogen interactions in a simple multicellular system.

1. How can I use this?

I can use C. elegans infection assays to compare how my resistant and non-resistant A. baumannii strains affect host survival or colonization. Worm mutants or RNAi lines can also help identify host pathways involved in responding to infection.

2. Why must I use this?

Cell culture alone cannot capture whole-organism level responses like innate immunity, stress signaling, or behavioral changes. C. elegans provides a simple but multicellular host with conserved pathways, making it a useful bridge between in vitro assays and more complex animal models. Using worms can help me evaluate whether my resistant strains show altered virulence or trigger different host responses, which is important for understanding the broader consequences of resistance evolution.

3. When will I use this?

I would use C. elegans models after generating resistant mutants or identifying bacterial genes that might influence virulence. Once I have strains I want to compare, worms allow me to quickly test infection outcomes, survival differences, or immune-related phenotypes. This would be especially helpful before moving into more resource-intensive mammalian systems.

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Powerful genetic engineering methods, such as transgenesis and gene targeting in mice, enable researchers to add, remove, or alter specific genes to examine their function within a live organism. This technology provides a precise and controlled method for examining the effects of biological treatments, such as probiotics or medications, on complex systems like cancer and immunology.

1. How can I use this?

I can create mouse models with specific genetic alterations related to immunological pathways, cytokines, or gut microbiome interactions through this technology. I can evaluate whether my next-generation probiotics actually improve chemotherapy outcomes, such as boosting immune cell activation, lowering inflammation, or increasing treatment sensitivity, by creating mice with modified immune responses or tumor microenvironments.

2. Why must I use this technology?

Unlike conventional animal models, transgenic and gene-targeted mice offer precise control over specific genes related to immunity, cancer progression, and host-microbe interactions. They aid in determining the precise mode of action of my probiotic strains and prevent findings that are confusing due to biological differences. Compared to other models, such as cell lines or wild-type animals, this technology provides me with a clearer and more mechanistic understanding of the process.

3. When will I use this technology?

This technique will be used after my in vitro data reveal that my probiotic candidates have promising immunomodulatory or chemo-enhancing properties. At that point, I need an in vivo system to evaluate the mechanism and ensure its safety and efficacy. When I want to test particular pathways, like removing a receptor that my bacteria are meant to stimulate, gene-targeted mice are particularly helpful. This is often the preclinical validation stage before proceeding to more advanced animal experiments.

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Although these immunology methods are not directly related to my next-generation probiotic project， but learning about them is still valuable for understanding advanced laboratory techniques and experimental design.

1. How can I use this?

Flow cytometry allows me to analyze thousands of individual cells within seconds by giving detailed information about their surface markers, internal proteins and overall functional state. Using fluorescent antibodies or dyes, I can identify which cells are activated and which belong to specific immune populations. Cell sorting extends this capability by physically separating selected cell types from a mixed sample. This lets me isolate pure populations of T cells, B cells, macrophages or other subsets for further experiments, such as gene expression analysis or functional assays. Together, these methods help me understand cellular diversity and immune responses with precision and speed.

2. Why must I use this?

These techniques are essential because the immune system is highly complex and bulk measurements often hide important differences between individual cells. Flow cytometry provides quantitative and single-cell resolution, which is critical for distinguishing subtle immune changes that may be missed in traditional assays. Cell sorting is equally important when I need clean and well defined cell populations to study specific mechanisms or to perform downstream experiments without contamination from unrelated cell types. Both methods are standard in immunology because they offer accuracy, scalability and deep insight into how immune systems function in health and disease.

3. When will I use this?

I will use flow cytometry when I need to measure immune cell composition, activation states, cytokine expression or cell cycle progression with high precision. It is especially valuable when studying complex samples, such as blood, lymphoid tissues or immune stimulated cultures. I will use cell sorting when I need to purify a specific immune subset for detailed analysis or controlled experimental manipulation. These methods are particularly important during experiments that require clear identification of immune phenotypes or functional validation of specific cell populations.

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1. How can I use this?

I can use High-throughput screening (HTS) to evaluate how different microbial strains, genetic constructs or metabolic pathways influence the production or regulation of specific metabolites. For my project, HTS can help me identify probiotic variants that show the strongest ability to produce beneficial metabolites or suppress harmful ones. It also allows me to test how engineered bacteria respond to various environmental conditions, nutrients or host-related factors. By using HTS, I can quickly narrow down large sets of candidates and focus on the most promising biological designs.

2. Why must I use this?

I must use HTS because metabolite modulation involves many interacting pathways and testing them one by one would be slow and inefficient. Engineering next-generation probiotics requires exploring large design spaces such as different promoters, enzymes, microbial chassis, metabolic regulators and growth conditions. HTS gives me a scalable way to examine all of these variables in parallel and detect subtle metabolic changes that may not be visible in small scale experiments. It also improves the reliability of my findings by generating robust datasets with strong statistical power. Without HTS, identifying the best engineered strains or the most influential metabolic pathways would take far longer and might even miss important combinations that only appear when screened at large scale.

3. When will I use this?

I will use HTS during the early and mid-development stages of my probiotic project, when I need to explore many design possibilities. On early stage, HTS helps me scan large libraries of engineered constructs to find which genetic designs have the strongest metabolic effects. As the project progresses, I can use HTS to test environmental factors such as pH, nutrients or host-derived signals that influence metabolite production. Later, before moving into animal studies, HTS can help me to confirm which engineered strains remain stable, efficient and responsive across a wide range of conditions. This lets me confidently select only the most promising candidates for more advanced testing.

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1. How can I use this?

Organoids allows me to work with miniaturized and three-dimensional tissue models that closely mimic real human organs. These systems can be used to study how cells grow, differentiate and respond to stress or mutations in a controlled environment. I can use organoids to examine how specific tissues behave under different metabolic conditions or how genetic alterations influence developmental pathways. They also give me a practical system to test potential interventions such as drugs, metabolites and engineered biological tools before taking the work into animal studies.

2. Why must I use this?

Organoids capture many structural and functional features of real organs, which makes them far more physiologically relevant than traditional two-dimensional cell culture. They allow the study of complex tissue behavior, including cell–cell interactions, metabolic gradients and developmental signaling pathways. For cancer and developmental biology, this is essential because disease mechanisms often depend on the spatial organization of cells and the microenvironment. Organoids offer a human-relevant platform for testing hypotheses while reducing reliance on in vivo models.

3. When will I use this?

I would use organoids when I need a system that sits between traditional cell lines and full animal models. This is particularly useful when studying early developmental processes, tumor initiation or tissue-specific metabolic regulation. In relation to my next-generation probiotic project, organoids may be useful only if I need to explore how microbial metabolites affect specific human tissues such as gut epithelium or liver tissue in a controlled setting. If my project focuses strictly on microbial engineering rather than tissue-level responses, organoids may be less directly relevant. However, if I want to test how engineered probiotics influence epithelial barrier function or host metabolic pathways, using gut or intestinal organoids could provide valuable and mechanistic insights before moving into animal studies.

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1. How can I use this?

I can use transgenesis and gene targeting in mice to study how specific genetic changes influence metabolism, gut physiology and host–microbe interactions. By generating mice with specific genetic modifications, I can clearly see how changes in certain genes influence metabolic pathways in the whole organism.These mouse models give me a controlled system to test how engineered probiotics interact with hosts that have known genetic backgrounds. This helps me understand which host genes or pathways are important for metabolite regulation.

2. Why must I use this?

I must use these techniques because metabolism is influenced by many tissues and signaling networks that cannot be replicated in cell culture alone. I need an live organism to truly understand how genetic changes to shape the gut environment, immune system and the composition of the microbiome. Transgenic and gene-targeted mice allow me to establish cause-effect relationships between host genes and metabolic outcomes. This is crucial for developing next-generation probiotics that must work consistently in the human gut, where host genetics strongly influence metabolic processes.

3. When will I use this?

I will use mouse models during the testing and optimization phases of my probiotic project. Early in development, I can use gene-targeted mice to identify which host pathways most strongly influence metabolite levels. Later, I can test engineered probiotics in transgenic mice with altered metabolic pathways to see whether the probiotics can correct or modulate those metabolites.Before I move toward translational studies, I will use these mouse models to make sure the engineered strains are safe, stable and actually work inside a living organism's bodies.

# data[].attributes.headline
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.subject
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

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1. How can I use this?

I can use C. elegans as a powerful model organism to study conserved biological processes such as development, aging, neurobiology, metabolism, and cell death. I can also apply extensive genetic tools such as mutants, RNA interference, and transgenics to manipulate gene function and analyze disease-related pathways. Because its genome shares 50–80% homology with humans, I can use worms to model human disease mechanisms in a simpler system.

2. Why must I use this?

I must use C. elegans because it provides a fast, inexpensive, and ethical organism for research. Worms grow quickly with a short life span of about 14–21 days, enabling rapid studies of aging and gene regulation. The lecture emphasizes that discoveries in worms have directly led to major breakthroughs in humans, such as identifying the programmed cell death pathway that is highly conserved and important in cancer biology. In addition, because worms reproduce without mating and require little space or cost to maintain, large-scale genetic screens become feasible, something difficult and expensive in mammals.

3. When will I use this?

I will use C. elegans when my research requires a genetic system to uncover gene function in a whole organism or to model diseases such as neurodegeneration, cancer, or metabolic disorders. I may also apply C. elegans in drug or RNAi screening to rapidly identify therapeutic targets before moving to mammalian systems.

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How can I use this?

I can utilize CRISPR/Cas9 gene editing or microinjection to generate transgenic C. elegans strains carrying fluorescent reporters that light up when the innate immune system is triggered. Instead of complex procedures with mice, I can simply infect these worms with bacteria on agar plates or in liquid culture. By using RNA interference by feeding, I can also systematically silence thousands of host genes to identify exactly which pathways are required to survive the bacterium induced oxidative stress and colonization.

Why must I use this?

C. elegans is essential because it allows for high-throughput, whole-organism screening that is impossible in mice, organoids, or cellular models. Because the worm is transparent, I can visually track the accumulation of bacteria in the worm and the simultaneous activation of immune reporters in a living host without dissection or complex imaging. This model isolates the innate immune response, allowing me to dissect the evolutionarily conserved innate defense mechanisms against bacteria without the confounding variables of the adaptive immune system.

When will I use this?

I will use the C. elegans model at the start of the discovery pipeline, specifically for high-volume screening of antimicrobial compounds or bacterial virulence factors. After identifying a library of potential drug candidates that rescue the worms from bacterial virulence, or pinning down specific bacterial genes that drive cellular stress responses, I will then select the most promising few candidates to validate in the more labor intensive murine models or complex organoid systems.

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How can I use this?

I can utilize gene targeting in embryonic stem cells to generate Upk1a-deficient mice, selectively inactivating the gene responsible for the urothelial receptor for FimH from Klebsiella pneumoniae. Instead of relying on cellular or molecular adhesion assays, I can challenge these mice with K. pneumoniae and perform longitudinal analysis of bacteriuria and tissue bacterial load. By comparing these knockouts to wild-type mice, I can directly test how the absence of the specific mannose-presenting scaffold alters the formation of Intracellular Bacterial Communities and the recruitment of neutrophils to the bladder mucosa in a living, physiologically complete system.

Why must I use this?

Transgenic (specifically gene-targeted) mice are essential because cell cultures and organoids cannot fully capture the systemic innate immune response and the complex hydrodynamics of micturition that define urinary tract pathogenicity. Klebsiella infections triggers a "scorched earth" defense called exfoliation, where infected cells detach to flush out bacteria; this process relies on a signaling cascade that requires the structural integrity of the whole bladder wall. The Upk1a knockout mouse reveals a critical paradox unavailable in 2D cultures: without the receptor, the host fails to sense the infection (blunted TLR4 signaling), leading to reduced bacterial clearance despite the lack of binding targets, which is a mechanism that requires a whole-organism model to dissect.

When will I use this?

I will use this mouse model when moving from identifying adhesion inhibitors to validating host-response mechanisms. After discovering potential anti-adhesive agents in vitro, I must test them in the Upk1a knockout model to distinguish between simple blocking of attachment and the complex modulation of the host's immune system. This allows me to determine if a therapy effectively clears the infection or merely masks it, preventing the immune activation necessary for long-term resolution before considering clinical translation.

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Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

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From this lecture, I learned that organoids give us a model that is much closer to real human physiology than 2D culture, but still easier and faster than using animals. Because they self-organize, keep multiple cell types, and maintain patient-specific features, I can use organoids to study diseases, test viruses, screen drugs, and even perform simple gene-editing experiments more reliably than with cell lines. These applications make organoids useful whenever I need a system that reflects human tissue behavior—such as modeling cystic fibrosis, microcephaly, tumor response, or checking drug toxicity. Organoids are indeed a practical choice when I need biological relevance without the complexity and time required for animal models.

# data[].attributes.headline
Luong Thi Minh Trang - 314302021 -   ltmtrang.ls14@nycu.edu.tw

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 -   ltmtrang.ls14@nycu.edu.tw

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How can I use this technique?

I can apply high-throughput screening when I need to test many variables (e.g., compounds, gene knockdowns) in my project quickly and in parallel. HTS allows me to evaluate multiple conditions under consistent experimental settings, which increases efficiency and comparability.

Why must I use this technique?

I must use HTS because manual, one-by-one testing is too slow and may miss hits. HTS brings automation, higher throughput, and better control over variables, helping to identify significant modulators of immune responses reliably.

When will I use this technique?

I will use HTS at the early discovery stage of the project, when screening many candidates (small molecules, antibodies, gene edits) for their effect on an immune outcome. Once hits are identified, I will switch to focused validation and mechanistic follow-up.

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How can I use this?

The tools we learned from C. elegans research can actually be applied directly to B cell metabolism. For example, RNAi lets me quickly knock down metabolic genes to see what changes, and CRISPR can create stable knockout B cell lines for deeper studies. I can also use genetic libraries to screen many genes at once and find the ones that really matter for metabolism. With techniques like flow cytometry and metabolic dyes, I can even track metabolic states at the single-cell level.

Why must I use this?

Because B cell metabolism is too complex to figure out by looking at one gene at a time. Genetic tools give me a systematic and unbiased way to pinpoint which genes truly drive metabolic changes, not just correlate with them. Plus, many metabolic pathways discovered in C. elegans are conserved in mammals, so these methods help me build solid, reproducible findings that are more likely to apply in real immune systems.

When will I use this?

I’ll use these methods throughout the whole research process. In the beginning, genetic screens help me generate hypotheses and narrow down key genes. In the middle stage, CRISPR or RNAi allows me to test which genes actually affect metabolism. Later, techniques like RNA-seq, ChIP-seq, or metabolomics help me figure out the mechanism behind those effects. And finally, in disease-related studies, I can use mouse models or primary human B cells to see whether these metabolic genes play roles in autoimmune disease, lymphoma, or infections.

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How can I use this? C. elegans can be used to study conserved biological processes such as cell fate determination, development, stress response, and neuronal signaling. The worm is transparent, genetically traceable, and has a completely mapped lineage, which allows quick analysis of gene function in a whole-animal context. C. elegans is mostly used to study neuronal development and behavior, cell lineage tracing, and aging research, which does not overlap greatly with my ovarian cancer focus; however, I could still use C. elegans as a simple system to explore basic gene functions or signaling pathways that are evolutionarily conserved before examining them in more complex models.

Why must I use this?  Many fundamental biological mechanisms, such as apoptosis, developmental signaling and cell-fate determination, were first discovered in C. elegans. Its short life cycle and relatively easy genetics make it ideal for quick functional screens or for studying pathways that would be difficult to analyze directly in mice. Worm studies can provide foundational mechanistic insights that might help interpret conserved processes in higher organisms.

When will I use this? I do not expect to use C. elegans in my current work, since my research relies heavily on mouse models and organoid cultures. However, it could be useful in the future for preliminary genetic screens to study conserved pathways at a basic level before validating them in mammalian systems.

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陳宥均 student ID 114101030 aidanchen38@gmail.com

# data[].attributes.subject
陳宥均 student ID 114101030 aidanchen38@gmail.com

# data[].attributes.body
1. How can I use this?

I can utilize Caenorhabditis elegans as a powerful platform for conducting rapid genetic screens and in vivo imaging. Practically, this involves employing techniques such as RNAi by simply feeding the worms bacteria that express dsRNA, allowing for easy and scalable gene knockdown without complex injection protocols. I can also take advantage of the organism's natural transparency to use fluorescent protein reporters to visualize protein location, cell migration, and organelle dynamics in a living, intact animal. Furthermore, I can use the detailed maps of its cell lineage to trace the developmental fate of single cell, manipulating pathways to observe how they alter the invariant developmental program.

2. Why must I use this?

It is essential for research that requires high-throughput analysis in a whole-organism context, which is often prohibitively expensive or time-consuming in mouse models. The worm's short life cycle of about three days and its large brood size allow me to generate statistically significant data on genetics and longevity in a fraction of the time required for vertebrate models. Additionally, because the wiring diagram of its nervous system is fully mapped, I must use this model if I want to understand how neural circuits drive behavior at a single-neuron resolution.

3. When will I use this?

I will use this model primarily when I need to dissect conserved biological pathways, such as apoptosis, aging, or insulin signaling, before validating them in mammalian systems. It is particularly useful when I am investigating the genetics of aging and longevity, as the worm's short lifespan allows for survival studies that would take years in mice to complete in just a few weeks. I will also turn to this model during the early discovery phase of a project to perform forward genetic screens—mutating the genome to find the genes responsible for a specific phenotype—thereby identifying novel genetic players that can later be studied in humans or other higher organisms.

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陳宥均 student ID 114101030 aidanchen38@gmail.com

# data[].attributes.subject
陳宥均 student ID 114101030 aidanchen38@gmail.com

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1. How can I use this?

I can use the knowledge gained from this lecture to design and generate sophisticated mouse models that manipulate specific sequences within the genome. Practically, this involves utilizing techniques such as homologous recombination or CRISPR-Cas9 to create knockout or knock-in mice, allowing me to remove a gene’s function or introduce a specific mutation. By understanding the breeding strategies required to maintain these lines, I can effectively manage colonies to produce the specific genotypes needed for experimentation, ensuring that the genetic background remains consistent and reliable for analysis.

2. Why must I use this?

I must use transgenic and gene-targeting approaches because in vitro cell culture models often fail to capture the systemic complexity of an entire organism. While cells in a dish can show molecular mechanisms, they cannot replicate the interactions between different organ systems, the immune response, or complex behavioral phenotypes. Mice share a high degree of genetic homology with humans, making them the gold standard for studying gene function in a physiological context.

3. When will I use this?

I will use these methods when I have identified a candidate gene or pathway in preliminary studies and need to validate its function in a living system. This often occurs after initial high-throughput screens or cell-based assays have highlighted a potential target, and the research moves into a pre-clinical phase. Specifically, I will rely on these techniques when studying developmental biology to observe how gene loss affects embryogenesis, or when modeling complex human diseases like cancer or autoimmunity that require a functional immune system and tissue microenvironment.

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陳宥均 student ID 114101030 aidanchen38@gmail.com

# data[].attributes.subject
陳宥均 student ID 114101030 aidanchen38@gmail.com

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1. How can I use this?

I can utilize the concepts from this lecture by establishing and manipulating advanced biological models that go beyond standard cell lines. Practically, this involves learning to culture three-dimensional organoids derived from pluripotent stem cells or patient tissues to mimic organ architecture. I can also employ gene-editing technologies, such as CRISPR-Cas9, within these complex models to introduce specific mutations and observe their effects on development or tumorigenesis in a spatially accurate context.

2. Why must I use this?

I must use next-generation modeling because traditional two-dimensional cell cultures often fail to predict clinical outcomes accurately. Standard cell lines grown on plastic lack the structural complexity, cellular heterogeneity, and crucial microenvironmental interactions, that exist in actual human tissues. Relying solely on older methods risks generating data that is reproducible in the lab but irrelevant to the patient. Therefore, adoption of these advanced models is essential to capture the true biological behavior of cancer and development.

3. When will I use this?

I will use these techniques primarily during the translational phase of research or when investigating complex developmental mechanisms that require tissue-level organization. Specifically, this becomes relevant when I need to validate a potential drug target in a physiologically relevant setting before moving to clinical trials. I will also deploy these models when studying personalized medicine, such as testing a battery of drugs on an individual patient’s tumor organoids to determine the most effective treatment strategy.

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陳宥均 student ID 114101030 aidanchen38@gmail.com

# data[].attributes.subject
陳宥均 student ID 114101030 aidanchen38@gmail.com

# data[].attributes.body
1. How can I use this?

The methods discussed in this lecture provide the technical framework for designing and executing large-scale biological experiments. I can implement Automation and utilize liquid handling robotics and automated readout systems to process thousands of samples efficiently, minimizing the human error associated with manual pipetting. Also, this method is great for analyzing big data, filtering out false positives and identifying genuine "hits."

2. Why must I use this?

High-throughput screening (HTS) is indispensable in modern biomedical research and drug discovery for several reasons:

Unbiased Discovery

Efficiency and Speed

Cost-Effectiveness

3. When will I use this?

I will likely encounter and utilize these methods during specific phases of research, for example, early-stage drug discovery: This is the primary application. I will use HTS at the very beginning of a project after a therapeutic target has been identified but before lead compounds exist. And also drug repurposing: If I am working on finding new uses for existing, FDA-approved drugs (screening a library of known drugs against a new disease model), I will use these screening methods to quickly identify potential treatments.

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How can I use this?

I can use transgenesis and gene targeting in mice to understand the molecular mechanisms by which probiotics reduce inflammation in colitis. By using knockout or conditional mouse models, I can test whether specific inflammatory genes such as iNOS, TNF-α, or IL-10 are required for the anti-colitic effects of my Lactobacillus strains. This will help me connect my in vitro findings (NO and cytokine suppression) with real biological outcomes in a living organism.

Why must I use this?

I must use genetically modified mouse models because in vitro cell culture systems cannot fully represent the complexity of immune regulation, gut barrier function, and host–microbe interactions in colitis. Gene targeting allows me to prove causation, not just correlation, by showing whether a specific gene is necessary or sufficient for disease progression or probiotic action. This is essential for producing scientifically credible and translationally relevant results.

When will I use this?

I will use transgenic and gene-targeted mouse models during the animal study phase of my Master’s research, after completing in vitro probiotic screening. At that time, I will move from cell-based assays to in vivo validation of probiotic efficacy and immune mechanisms in colitis.

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313303017 - Pham Nhat Phuong Trinh - pnhatptrinh@gmail.com

# data[].attributes.subject
313303017 - Pham Nhat Phuong Trinh - pnhatptrinh@gmail.com

# data[].attributes.body
How can I use this?

I can use these mouse-genetics methods to change a gene, add a reporter, or control when a gene is turned on or off. Techniques like transgenesis, CRISPR, or Cre/loxP let me test the function of specific sequences or regulatory elements directly in a living organism, instead of only relying on cell culture results.

Why can I use this?

I can use them because mice allow precise and stable genetic manipulation, and their physiology is close enough to humans for disease studies. These systems also give me control over timing and cell-type specificity, which is important when a full knockout or early misexpression would be lethal or misleading.

When can I use this?

I will use these tools when my project needs in vivo confirmation. For example, when I want to check whether a regulatory variant truly changes gene expression in the brain, or when I need to delete or activate a gene specifically in striatal neurons to see how it affects pathways relevant to HD. This is the stage where cell models are no longer enough and I need evidence from an intact neural system.

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Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

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Flow cytometry is valuable to me because it gives single-cell information that bulk assays can’t provide. I can use it to follow how different immune cell subsets change, whether they activate, proliferate, undergo apoptosis, or express certain cytokines, and even sort out the exact population I need.

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Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.tw

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When I learned biology in high school, I only imagined genetics in simple Mendelian rules, so I never thought about how complex real gene–gene interactions could be. This lecture made me realize that biological traits arise from dense interaction networks, not single genes, and that these relationships can be far more complex.

What impressed me most is that modern combinatorial and multiplexed screening actually provides a practical way to uncover these hidden interactions at a scale impossible to reason about intuitively. It made me realize that if we want to understand complex diseases or find new therapeutic strategies, we really need these high-throughput genetic methods, because they're the tools that can actually handle the complexity of real biology.

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Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.vn

# data[].attributes.subject
Luong Thi Minh Trang - 314302021 - ltmtrang.ls14@nycu.edu.vn

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The application I'm most interested in is structure-based drug design using X-ray crystallography, because it allows me to see the atomic details of a drug-binding pocket, as illustrated by the electron-density maps and the examples of transition-state analog inhibitors (e.g., Enalapril, Aliskiren). I realize that many successful drugs today were developed precisely because we could visualize enzyme active sites at atomic resolution.

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How can I use this?

I can use this system by creating floxed mice for genes like G6PD, TALDO1, or TKT, and crossing them with CD19-Cre or AID-Cre mice to make B cell-specific knockouts. Then I isolate their B cells and compare their development, antibody responses, and metabolic activity. If needed, I can add immunization (like NP-KLH), metabolic tracing, RNA-seq, or flow cytometry to study how the deleted gene changes B cell function in vivo.

Why must I use this?

I need this approach because whole-body knockouts often cause early death or non-B cell effects, making results unclear. Using a B cell-specific deletion lets me see the direct role of the gene in B cells without interference from other tissues. It also gives more reliable in vivo data than in vitro CRISPR or shRNA. This helps me understand how pathways like the PPP support B cell activation, antibody production, and disease processes.

When will I use this?

I will use this system when I want to study the function of a metabolic gene specifically in B cells—such as during immunization, infection, tumor growth, or autoimmune conditions. It is also useful when exploring how gene loss affects antibody responses or when searching for metabolic targets for future therapies.

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Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.subject
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.body
1. How can I use this?

I can use mouse transgenesis and gene targeting technologies to explore the function of specific genes in a living organism. By introducing a foreign gene through pronuclear injection or modifying endogenous genes using CRISPR/Cas9 or embryonic stem cell–based homologous recombination. I can generate mice that model human diseases such as cystic fibrosis, Alzheimer’s disease, cancer, or diabetes. These genetically engineered mice allow me to test therapeutic strategies, track cells in vivo through reporters like GFP, and control when and where genes are expressed using inducible (Tet-on/Tet-off) or conditional (Cre/loxP) systems.

2. Why must I use this?

I must use transgenesis and gene targeting because mice share ~95% of their genes with humans, making them the most accurate mammalian model system for ethically studying human biology and diseases. These techniques allow both loss-of-function and gain-of-function studies to reveal gene necessity and sufficiency in development, immunity, and pathology. Conditional and inducible systems are essential when gene deletion causes early lethality or different effects in different tissues, ensuring we can dissect the role of a gene in specific organs or time points.

3. When will I use this?

If I work on tumor suppressor genes, I may generate a conditional knockout mouse to study cancer development specifically in immune cells. In translational and clinical research, gene-edited animal models are indispensable for therapeutic testing and for generating xenotransplantation-compatible donor organs, such as genetically modified pigs for heart transplantation.

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Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

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Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.body
1. How can I use this?

I can use organoid technology to model human organs and diseases in vitro because organoids are 3D mini-organs that self-organize from stem cells and recreate key structural and functional features of the real tissues. I can grow intestinal, airway, brain, liver, tumor, or patient-derived organoids to explore cell differentiation, developmental biology, tissue regeneration, infection responses, and cancer progression. Organoids allow me to test drugs, perform genetic modification, and integrate technologies like single-cell sequencing to understand cell diversity and signaling pathways more precisely.

2. Why must I use this?

I must use organoids because they provide a human-relevant model system that overcomes limitations of 2D cell lines and animal models, which may not accurately represent human physiology or disease. They are essential tools for discovering personalized therapies, such as testing responses of cystic fibrosis or cancer patient–derived organoids to drugs before treatment. Since organoids can be genetically engineered and scaled up for screening, they are increasingly considered the next-generation system for clinical and translational research, helping to bridge the gap between lab findings and patient treatment outcomes.

3. When will I use this?

I will use organoids whenever my research requires a model that closely reflects human biology, especially in fields like cancer biology, developmental biology, neuroscience, immunology, or infectious disease. For example, I might use patient-specific organoids to study tumor heterogeneity, identify drug-resistant clones, or design targeted therapies.

# data[].attributes.headline
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.subject
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.body
1. How can I use this?

I can use High-Throughput Screening (HTS) to rapidly test thousands of chemical compounds, drugs, or biological molecules against a biological target in order to identify those with desirable effects. HTS integrates robotics, automated liquid handling, miniaturized assays, and powerful data analytics to quickly generate meaningful results. I can design assays to measure outcomes such as enzyme inhibition, cell viability, viral infection, immune signaling, or gene expression. This technique allows me to efficiently discover hit molecules, validate therapeutic targets, and explore genotype-phenotype relationships in disease-related pathways.

2. Why must I use this?

I must use HTS because modern biomedical challenges—like developing cancer therapies, antivirals, and personalized medicine strategies—require the ability to evaluate a massive number of potential drug candidates quickly and efficiently. Traditional assays are too slow and costly for large-scale drug discovery. HTS helps identify early-stage leads, eliminate ineffective compounds rapidly, and reduce research time and cost. It also enables the discovery of molecules that modulate complex biological systems, such as immune checkpoints or metabolic pathways, which is essential for advancing new treatments for diseases.

3. When will I use this?

I will use HTS whenever the goal of my research involves finding new therapeutic compounds, studying molecular interactions, or screening genetic modifications across large libraries.

# data[].attributes.headline
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.subject
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.body
1. How can I use this?

I can use flow cytometry to analyze individual cells in a mixed population by measuring their size, surface markers, protein expression, cell cycle stage, proliferation, apoptosis, and functional characteristics, all at high speed and with high precision. The lecture shows that flow cytometry can evaluate up to millions of cells per minute and detect multiple biological parameters simultaneously (e.g., CD markers, cytokines, phosphorylated proteins). I can use this technology to determine immune cell subsets (like CD4 or CD8 T cells), track cell division using dyes like CFSE, study cell signaling via phospho-flow staining, and even sort specific cell populations for further experiments.

2. Why must I use this?

Flow cytometry is a critical technique for modern biomedical research because it allows me to connect cell phenotype with function. For example, I must use it when I need to distinguish live and dead cells during drug testing, measure cytokine responses to infections, or identify early apoptosis by detecting phosphatidylserine exposure using Annexin V. It is one of the only methods that can provide multidimensional immunophenotyping, which is essential for studying diseases like cancer, autoimmune disorders, and immunodeficiency.

3. When will I use this?

I will use flow cytometry whenever my research requires identifying or characterizing specific cell types in a heterogeneous sample, such as blood, tumor tissue, bone marrow, or cell cultures. This includes projects involving immune cell profiling, vaccine studies, stem cell development, tumor immunology, or evaluating treatment responses.

# data[].attributes.headline
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.subject
Bui Truc Vy - 313302028 - vybui.oralie18@gmail.com

# data[].attributes.body
1. How can I use this?

This lecture teaches how complex traits arise from interactions between multiple genes, such as epistasis and synthetic lethality, instead of simple Mendelian genetics. I can use this knowledge to understand disease mechanisms, especially cancer, where parallel pathways and regulatory networks interact to determine cell fate. The lecture also introduces model organisms (like yeast) that allow high-throughput genetic screening and the discovery of global genetic interaction networks, which help map essential cellular functions. Additionally, principles from synthetic biology, such as toggle switches and the lactose operon as logic gates, can be applied to design genetic circuits, biosensors, or therapeutic gene networks.

2. Why must I use this?

I must apply these genetic interaction concepts because human diseases are complex, involving many genes working in networks rather than individually. Synthetic lethality, for example, provides a rationale for targeted cancer therapy—inhibiting two genes simultaneously can selectively kill cancer cells that rely on compensatory pathways.

3. When will I use this?

I will use this knowledge whenever I analyze disease genetics, drug response, or conduct research involving cellular pathways. Even in basic research, understanding gene-gene interaction networks and model organism systems guides experimental design and helps translate findings from yeast or bacteria to humans.

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In this week’s class, we learned about transgenesis and gene targeting in mice, including classical gene knockout methods and newer genome-editing approaches like CRISPR.

1. How can I use this?

I can use genetically modified mice to study how specific host genes influence susceptibility to A. baumannii infection. Knockout or CRISPR-edited mice would allow me to test how changes in immune pathways affect the response to resistant versus non-resistant strains.

2. Why must I use this?

Mouse models provide whole-organism context that cell cultures can’t. Using targeted gene modifications helps isolate the role of individual host genes in inflammation, bacterial clearance, or antibiotic response—factors that are crucial for understanding resistance mechanisms.

3. When will I use this?

I would use these models after identifying host pathways that seem important from in vitro experiments. Once I have candidate genes, transgenic or knockout mice can help confirm their roles during infection and compare how different A. baumannii strains behave in vivo.

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How can I use this?
I can utilize transgenesis and gene targeting to create mouse lines that either lack key immune-related genes (such as CCR2) or overexpress specific receptors involved in tau sensing. This lets me directly test how these genes shape monocyte behavior and tau spreading in the brain.

Why must I use this?
Because tauopathy is a complex in vivo process, and cell culture alone can’t capture the full immune–brain interaction. Genetically modified mice provide me with precise and controlled methods to determine whether a gene is truly required—or sufficient—for driving inflammation or tau transmission.

When will I use this?
I’ll use these models once I’ve identified promising targets from RNA‑Seq or in vitro assays. These mice serve as the backbone of my in vivo experiments, particularly when comparing disease progression between wild-type, knockout, and inducible gene-controlled lines.

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How can I use this?
I can use brain organoids or immune–organoid co‑culture systems to recreate a more realistic microenvironment for studying how tau interacts with monocytes. This lets me test tau uptake, inflammatory signaling, and cell–cell communication in a setting that’s much closer to human biology than standard 2D cultures.

Why must I use this?
Because tauopathy is a complex, multicellular process, and simple cell lines can’t capture the architecture, signaling gradients, or immune–neuron interactions that happen in vivo. Organoid-based models provide a more accurate platform for validating mechanisms before moving into animal studies.

When will I use this?
I’ll use these models after initial in vitro assays—once I’ve identified key pathways or genes from monocyte responses. Organoids serve as an intermediate step, where I test whether those findings still hold in a more physiologically relevant system before committing to mouse experiments.

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How can I use this?
I can utilize high-throughput screening methods to rapidly test large panels of small molecules, inhibitors, or genetic perturbations to identify those that affect monocyte activation, tau uptake, or tau-induced inflammation.

Why must I use this?
Because tau‑driven immune responses involve many pathways, guessing one target at a time is slow. High‑throughput screening lets me rapidly narrow down which pathways or compounds are actually worth following up in vivo.

When will I use this?
I’ll use it early in the project—right after identifying candidate genes or signaling pathways—to screen for hits in cell‑based assays before moving the most promising ones into mouse experiments.

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How can I use this?
I can use flow cytometry and cell sorting to identify and isolate specific immune cell populations—like infiltrating monocytes, microglia, or T cells—from mouse brains after tau injection, based on surface markers and activation status.

Why must I use this?
Because I need clean, well-defined cell populations to really understand who is responding to tau and how. Bulk tissue data can’t tell me which cell type is changing; flow and sorting let me link specific immune subsets to cytokine profiles, RNA-Seq data, or functional assays.

When will I use this?
I’ll use it after in vivo experiments—at key time points after tau or vector injection—to analyze immune infiltration, activation, and to sort monocytes or microglia for downstream RNA, protein, or ex vivo functional studies.

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How can I use this?

I can use combinatorial and multiplexed genetic screening to map gene–gene interactions, find epistasis, identify synthetic lethal pairs, and understand complex traits that don’t follow simple Mendelian rules. It’s essentially a toolkit for understanding how genes interact rather than acting independently.

Why must I use this?

Because most real biological systems—metabolism, signaling, disease pathways—are not controlled by single genes. They’re networks. If I only study one gene at a time, I miss the real biology. These strategies let you uncover hidden interactions, parallel pathways, and functional modules that traditional genetics can’t reveal.

When will I use this?

I’ll use it whenever you need to understand complex phenotypes, such as drug resistance, cancer metabolism (like the Warburg effect), aging circuits, or regulatory networks (toggle switches, repressilators). It’s also essential when designing synthetic biology circuits or doing large‑scale functional genomics.

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Angie Herold (廖愛穎, 313304026) - Herolda2002@gmail.com

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Angie Herold (廖愛穎, 313304026) - Herolda2002@gmail.com

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How can I use this? I can use transgenesis and gene-targeting to generate mouse models that allow us to study gene function in ovarian and fallopian-derived tumors. We breed mouse lines that use Cre/loxP or Tet-regulated Cre systems to induce tissue-specific and, in some cases, time-specific gene knockouts. This enables deletion of target genes only within the reproductive tract epithelium or in Müllerian lineage-derived tissues. I can use these mice to establish organoid cultures for subsequent analyses.

Why must I use this?  This is important because in vitro studies alone are often not sufficient to investigate cancer-related genes. Conditional knockout systems enable me to control where and when a gene is deleted, thereby helping to avoid early lethality or developmental defects. Temporal controls such as the Tet-O-Cre are also valuable when gene deletion should occur at a later life stage. These models better reflect in vivo disease mechanisms and provide organoids with the correct genetic background, making downstream assays (e.g., siRNA knockdown, RNA-seq, drug testing,…) more physiologically relevant.

When will I use this? I already work with my own Cre/loxP mouse colonies, and they are a central part of my current research. These include both constitutive (Amhr2-Cre) and inducible systems (Pax8-tetO-Cre and K8-Cre). They are necessary for us to generate tumor-derived organoids that serve as the basis for our later analyses.

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In this week’s class, we were introduced to organoid systems and how they can model human tissues in a way that better reflects in vivo physiology. I was particularly interested in how organoids can be used to study infection and drug responses, which made me think about their potential connection to my research on Acinetobacter baumanniiantimicrobial resistance.

1. How can I use this?

I can use organoid-based infection models—such as airway, lung, or intestinal organoids—to study how A. baumanniiinteracts with human tissues under more realistic physiological conditions. These models would allow me to examine epithelial damage, inflammatory responses, cytokine secretion, or cell death patterns caused by resistant versus non-resistant strains. Organoids also provide a controlled system for testing antibiotic efficacy or potential adjuvant therapies directly on human-like tissue structures.

2. Why must I use this?

Standard cell lines often oversimplify host–pathogen interactions, which can limit understanding of clinically relevant resistance mechanisms. Organoids offer better tissue architecture, multiple cell types, and more accurate signaling environments. This makes them valuable for capturing the complex ways in which A. baumannii affects human tissues, especially when evaluating virulence factors or antibiotic responses of resistant mutants. Using organoids can help generate data that more closely reflects how infections behave in real patients.

3. When will I use this?

I would consider using organoids after identifying specific resistance-associated mutations or phenotypes from my bacterial evolution experiments. Once I have resistant and sensitive strains to compare, organoids would be useful for testing whether these variants differ in their ability to invade tissue, trigger inflammation, or withstand antibiotic treatment in a more realistic model. They would also be helpful if I aim to explore potential therapeutic strategies or host-directed treatments.

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In this week’s class, we focused on the concept of high-throughput screening (HTS) and how it can be used to study complex biological questions. What stood out to me is how HTS makes it possible to test thousands of conditions at once and quickly narrow down key molecules or genetic factors. It also helped me think about how this approach could be applied to my own research on Acinetobacter baumannii antimicrobial resistance.

1. How can I use this?
I can use high-throughput screening (HTS) to systematically identify compounds or genetic factors that influence A. baumannii survival or antibiotic response. By designing either a target-based assay—such as monitoring efflux pump activity—or a phenotype-based assay like viability under tigecycline or imipenem exposure, I can screen large chemical or genetic libraries in parallel. After validating the assay with good controls and a strong Z-factor, automation allows me to test thousands of samples efficiently and select promising hits for follow-up studies.

2. Why must I use this?
HTS provides a scalable and quantitative framework that helps reveal important modulators of antibiotic resistance that small-scale experiments might miss. It improves reproducibility through automation, supports both biochemical and cell-based readouts, and accommodates many types of libraries. This makes it especially useful for identifying novel drug candidates or discovering pathways involved in heterogeneous or spontaneous resistance in A. baumannii.

3. When will I use this?
I would use HTS when I need to screen large compound or genetic libraries to find potential inhibitors of resistance pathways, or when exploring genetic interactions that contribute to resistance development. It will also be helpful after generating resistant mutants so I can test their responses to many antibiotics or stress conditions at once. High-content imaging or phenotypic assays can further provide multiparameter data on bacterial behavior or host–pathogen interactions.

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NGUYEN THI MINH HANG - 313302027 - minhhangng28@gmail.com

1. How can I use this?

I can use organoid cultures to study tumors ex vivo by growing 3D structures derived from tumor tissue. Because organoids preserve many architectural and functional features of the original tumor, I can use them to analyze gene function, examine tumor heterogeneity, assess pathway activity, and test drug responses in a system that more closely reflects in vivo conditions than traditional 2D cell lines.

2. Why must I use this?

I must use organoids because they better replicate the complexity of real tumors, including cellular diversity, structural organization, and patient-specific genetic backgrounds. Since cancer is highly heterogeneous, organoids provide more accurate and reliable platforms for evaluating gene perturbations, pathway changes, and therapeutic responses compared to standard cell-line models. Their ability to be expanded long-term also supports reproducible molecular and functional assays.

3. When will I use this?

I will use organoids whenever I need a model that closely mimics in vivo tumor biology. They are valuable for experiments such as gene knockdown, transcriptomic profiling, functional assays, and drug testing. As a core system in my research, organoids serve as the foundation for many of my studies using mouse-derived tumor samples.

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NGUYEN THI MINH HANG - 313302027 - minhhangng28@gmail.com

1. How can I use this?

I can use high-throughput screening to systematically identify genes, pathways, or small molecules that regulate cellular metabolism by combining CRISPR libraries or compound libraries with metabolic assays such as glycolytic rate, mitochondrial function, or cell viability under stress. This allows me to rapidly uncover metabolic dependencies in a way that targeted, low-throughput experiments cannot.

2. Why must I use this?

I must use HTS because cellular metabolism is controlled by complex and interconnected networks. Low-throughput methods cannot capture the full range of regulators involved. HTS provides an unbiased, large-scale approach to discover unexpected metabolic factors, prioritize meaningful targets, and efficiently support mechanistic research or therapeutic development.

3. When will I use this?

I will use HTS when defining the metabolic features of different cell types, when screening genes or compounds that influence cellular function, and when validating candidate pathways or testing drug combinations. HTS is also valuable for studying how metabolism changes during activation, differentiation, disease progression, or treatment responses, especially when combined with single-cell platforms.

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NGUYEN THI MINH HANG - 313302027 - minhhangng28@gmail.com

1. How can I use this?

I can use flow cytometry to measure the physical and chemical properties of thousands of cells per second as they pass through a laser. By labeling cells with fluorescent markers, I can identify and quantify different cell types in a mixed sample. With FACS, I can also sort and isolate specific cell populations with high purity for further experiments.

2. Why must I use this?

I must use this because it provides rapid, quantitative, single-cell analysis that bulk methods cannot offer. Flow cytometry is essential for detecting abnormal cell populations in diseases like leukemia, monitoring immune responses, and studying rare but important cell subsets. It gives detailed, multi-parameter information necessary for both clinical diagnostics and research.

3. When will I use this?

I will use this whenever I need to identify, count, or separate specific cell types from complex samples. This applies in immunology, cancer diagnostics, stem cell tracking, and basic research where pure cell populations are required to study their functions.

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NGUYEN THI MINH HANG - 313302027 - minhhangng28@gmail.com

1. How can I use this?

I can use combinatorial and multiplexed genetic techniques to test many gene interactions at once by creating multiple genetic variants or synthetic circuits. These methods let me observe how different genes affect each other and help build functional maps of genetic networks in models like yeast or engineered cells.

2. Why must I use this?

I need to use these approaches because complex traits and diseases depend on interactions among many genes. They help identify essential genes, uncover therapeutic targets, and clarify how biological pathways work together. They also support synthetic biology by enabling the design of genetic circuits with new functions.

3. When will I use this?

These techniques are used in research on complex diseases, in high-throughput gene screens, and in developing targeted therapies. They are also applied in synthetic biology projects that require constructing artificial regulatory circuits or studying network-level cellular functions.

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Laishram Yashmine Devi, 313302024, yoonseri121@gmail.com 

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Laishram Yashmine Devi, 313302024, yoonseri121@gmail.com 

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1. How can I use this?

I can use intestinal organoids as an ex vivo model to test how my probiotic candidates influence epithelial health, inflammation, and barrier function. Their 3D structure and multiple cell types allow me to study host–microbe interactions in a system that closely resembles the human gut.

2. Why must I use this?

I must use organoids because traditional 2D cell lines cannot fully replicate the architecture or cellular diversity of the intestine. Organoids provide a more accurate and translational model, helping me better predict whether my Lactobacillus strains will have protective or therapeutic effects in colitis before moving to animal studies.

3. When will I use this?

I will use organoid experiments after completing basic in vitro screening assays (acid/bile tolerance, Caco-2 adhesion). This will serve as a functional evaluation step before beginning my in vivo mouse colitis experiments.

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313303017 - Pham Nhat Phuong Trinh - pnhatptrinh@gmail.com

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313303017 - Pham Nhat Phuong Trinh - pnhatptrinh@gmail.com

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How can I use this?

I can use organoids as a three-dimensional, physiologically relevant human model to study development, disease mechanisms, and therapeutic responses. By differentiating pluripotent stem cells into brain, gut, I can recreate layered architecture and multicellular interactions that are impossible to capture in 2D culture. In practice, I can apply organoids to model neurodevelopmental defects, perform genetic manipulation at the iPSC or organoid stage, combine them with single-cell sequencing to validate cellular composition, and use them for drug testing or toxicity screening in a more realistic tissue environment.

Why can I use this?

I can use organoids because they are human-derived and capable of capturing key cellular interactions that 2D cultures cannot. It offers strong advantages such as human physiological relevance, scalability once established, compatibility with modern genome engineering tools, and the ability to generate personalized disease models from patient-derived iPSCs. These features make organoids a suitable model when my research requires a higher level of biological complexity than conventional 2D systems.

When will I use this?

I will use organoids when my research question requires tissue-level organization or multicellular interactions. These contexts benefit from organoids’ ability to mimic aspects of human brain architecture that are absent in 2D culture.

However, after discussing with my PI, I also realized that organoids are not the optimal choice, especially for Huntington’s disease research. Although recent protocols can generate cortical or striatal-like organoids, these 3D models still do not reproduce the full architecture, signaling environment, or connectivity of the human brain, and lack standardized protocols. In addition, organoid culture requires specialized matrices, long differentiation timelines, and higher consumable usage, making it significantly more expensive than maintaining 2D cultures. These limitations make organoids far less reproducible than 2D iPSC-derived neuronal cultures, which routinely achieve high consistency. Nevertheless, I still keep in mind that this is an early stage of 3D organoid technology. As protocols become more standardized and costs progressively decrease, these models will likely mature into a far more reliable and powerful system that can eventually mimic human physiology with much greater fidelity.

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1. How can I use this?

Organoids are three-dimensional structures derived from stem cells that reproduce the architecture and functions of human organs. They can be used to model development, to study genetic or infectious diseases, to test drugs, and to explore regenerative therapy strategies. Their culture makes it possible to generate mini-organs such as intestine, brain, liver, or lungs, offering a relevant and manipulable experimental model.

2. Why must I use this?

This method is essential because it offers a faithful representation of human physiology, much more than 2D cell cultures or animal models. Organoids make it possible to understand pathological mechanisms, to develop personalized medicine approaches, and to reduce dependence on animal models. They provide a robust platform for drug screening, tissue repair, and the study of cellular diversity.

3. When will I use this?

Organoids are mobilized whenever it is necessary to reproduce the complexity of human tissue in vitro. They are used in fundamental research on development, in modeling diseases such as cystic fibrosis or cancer, in studies of viral infections (such as Zika or SARS-CoV-2), and in regenerative medicine projects. They become essential whenever an experimental approach must combine physiological relevance with the possibility of genetic or pharmacological manipulation.

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1. How can I use this?
Methods in Immunology such as Flow cytometry makes it possible to analyze several cellular parameters simultaneously at the individual scale. It relies on the detection of scattered light and fluorescence emitted by specific probes. It makes it possible to phenotype cells, to measure protein expression, to follow proliferation, to analyze the cell cycle, or to detect apoptosis. It can also be used to sort cells (FACS), to perform multiplexing, or fluorescent barcoding.

2. Why must I use this?
This method is essential to characterize complex cell populations quickly and precisely. It makes it possible to obtain robust quantitative data, to detect rare events, and to analyze dynamic processes. It is indispensable in immunology, oncology, cell biology, and experimental medicine. It offers unmatched multiparametric resolution for the study of cells in suspension

3. When will I use this?
It is used in immunological analyses, signaling studies, treatment monitoring, or biomarker screening. It is applied in stimulation protocols, intracellular staining, or cell sorting. It is mobilized whenever fine, rapid, and multiparametric cell characterization is required.

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1. How can I use this?

Combinatorial and multiplexed genetic strategies make it possible to test many gene interactions simultaneously using high-throughput screening approaches. They rely on the creation of synthetic genetic networks, the study of epistatic interactions, or the analysis of phenomena such as synthetic lethality. These methods use models such as yeast or artificial genetic circuits to map functional networks and to understand the logic of interactions.

2. Why must I use this?

These approaches are essential to decipher the complexity of human diseases, which often depend on multiple interactions between genes and metabolic pathways. They make it possible to identify essential genes, to reveal therapeutic targets, and to understand the structure of biological networks. They also provide a basis for synthetic biology, making it possible to design artificial genetic circuits capable of new functions such as cellular memory or oscillators.

3. When will I use this?

These strategies are used in research projects on complex diseases, in screening genes involved in cancer or metabolic diseases, and in designing targeted therapies based on genetic interactions. They are also involved in synthetic biology to create artificial regulatory circuits, and in network studies to map cellular functions. They are mobilized whenever it is necessary to explore or manipulate genetic complexity beyond Mendelian models.

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1. How can I use this?
Protein-DNA interactions can be studied by in vitro methods such as EMSA, DNase I footprinting, or SELEX, and by in vivo approaches such as ChIP, ChIP-seq, or ChIP-on-chip. Biophysical techniques such as SPR or ITC make it possible to measure affinity and dissociation constants. These tools make it possible to identify target sequences, to map binding sites, and to quantify the strength and dynamics of interactions.

2. Why must I use this?
This makes it possible to understand how a protein interacts with DNA, which is essential to decipher transcriptional regulation, to identify genomic targets, and to explore epigenetic mechanisms. These methods make it possible to characterize regulatory networks, to develop diagnostic tools, and to design targeted therapies. They offer fine and quantitative resolution of molecular mechanisms.

3. When will I use this?
These techniques are used in research projects on transcription factors, repair enzymes, or regulatory proteins. They are involved in studies of cell differentiation, tumor transformation, or immune response. They are mobilized whenever the function of a DNA-binding protein must be analyzed.

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1. How can I use this?

RNA technologies bring together a set of methods that make it possible to detect, analyze, and manipulate RNA molecules. They include classical techniques such as Northern blot, EMSA, RT-PCR, and RNA-FISH, as well as modern approaches like RNA-Seq, RNA interference (RNAi), RNA vaccines, and CRISPR systems targeting RNA. These tools make it possible to study the expression, stability, localization, and interactions of RNA in cells.

2. Why must I use this?

These methods are indispensable to understand post-transcriptional regulation, the dynamics of gene expression, and the mechanisms of RNA degradation. They offer powerful means to identify biomarkers, to develop innovative RNA-based therapies, and to explore genetic correction strategies. They are also crucial for the design of modern vaccines and targeted treatments against genetic or infectious diseases.

3. When will I use this?

RNA technologies are used in research projects on gene expression regulation, in transcriptomic studies, and in the development of new therapies. They are involved in viral RNA detection, vaccine design, expression profiling by RNA-Seq, or treatments using RNAi. They are mobilized whenever it is necessary to explore the role of RNA as a central molecule of biology and medicine.

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1.How can I use this?
Molecular cloning makes it possible to modify DNA in order to study or exploit the expression of a gene. This method is used to insert a sequence into a vector, to mutate it, to amplify it, or to delete it. It relies on techniques such as enzymatic digestion, ligation, PCR, or more advanced approaches like Gibson assembly or Infusion cloning. Once the gene is cloned, it can be transferred into a host cell by chemical or viral methods in order to observe its biological effects or to produce the corresponding protein.

2.Why must I use this?
This method is indispensable to understand the function of a gene, to validate bioinformatic hypotheses, to produce recombinant proteins, or to develop therapeutic vectors. It makes it possible to introduce targeted mutations, to create cellular or animal models, and to test biological mechanisms. Without cloning, it is impossible to manipulate gene expression in a controlled and reproducible way.

3. When will I use this?
It is used whenever gene expressions must be modified in a biological system. Cloning is involved in functional studies, protein production, validation of therapeutic targets, or the creation of experimental models. It is also employed in laboratories of molecular biology, genetics, virology, or translational medicine.

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Elise, Student ID: 114350804 

 elise.gascher.bt14@nycu.tw.edu 

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Elise, Student ID: 114350804 

 elise.gascher.bt14@nycu.tw.edu 

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• How can I use this?

I can use organoid systems to model human tissues and diseases in a controlled, three-dimensional environment. By deriving organoids from adult stem cells or pluripotent stem cells, I can recreate key structural and functional features of organs. This allows me to study development, genetic diseases, cancer, infections, and to perform drug testing or toxicity assays. Organoids can also be combined with single-cell sequencing, organ-on-chip platforms, and genetic engineering tools for deeper mechanistic studies.

• Why must I use this?

I should use organoids because they represent human physiology more accurately than traditional 2D cell lines or some animal models. They are fast to establish, genetically tractable, scalable for drug or genomic screens, and can be personalized from patient-derived cells. Organoids enable disease modeling that captures human-specific features—such as CFTR defects in cystic fibrosis or neural progenitor vulnerability to Zika—while reducing ethical and experimental limitations of in vivo systems.

• When will I use this?

I will use organoids when I need human-relevant models for development, disease mechanisms, or therapeutic discovery. This includes studying cancer evolution, testing drug responses, modeling genetic disorders, analyzing host–pathogen interactions, or investigating tissue development such as cerebral organoids for microcephaly studies. I will also use organoids for personalized medicine, toxicity assessments, and high-throughput or high-content screens that require tissue-like complexity.

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How can I use this?

To study B cell metabolism, I would use germinal center (GC) organoids, because they can recreate a 3D lymphoid environment that normal 2D cultures cannot provide. Experimentally, I would embed B cells in a 3D hydrogel matrix and include supporting cells that express CD40L and BAFF, allowing the organoids to maintain GC-like activity for several days. After establishing the system, I would analyze metabolic changes using flow cytometry for glycolysis- or OXPHOS-related markers and use LC-MS or GC-MS to measure key metabolites. This setup would help me observe how B cell activation, class switching, and differentiation are linked to specific metabolic pathways.

Why must I use this?

I would choose organoids because 2D cultures cannot accurately maintain germinal center B cell metabolism. GC B cells are short-lived and strongly influenced by spatial organization (such as differences between dark-zone and light-zone B cells) which only a 3D system can model. Organoids also allow more reliable analysis of metabolic transitions during B cell activation and plasma cell differentiation, since they generate enough cells for metabolomics and single-cell profiling.

When will I use this?

I would use organoids when I need to explore how B cell metabolism shifts during activation and differentiation, because organoids can generate GC-like responses in just a few days. They are also useful when screening multiple metabolic inhibitors or pathway modulators, since the high-throughput format allows many conditions to be tested at once. In later stages of research—such as identifying metabolic biomarkers, comparing donor-specific metabolic differences, or evaluating vaccine or immunotherapy responses—organoids remain valuable because they closely reflect human biology.

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Organoid technology is a next-generation 3D culturing technique that makes it possible for human tissues, such as tumors or healthy epithelial cells, to develop in a physiologically accurate, miniature form. Organoids provide an effective platform for determining complex interactions between cancer cells, immune cells, and microbiological products because they closely resemble the shape, function, and genetics of actual human tissues. This makes the technology highly valuable in my study, as it allows me to test how my probiotic strains influence chemotherapy response in a human-like environment before moving on to an animal model.

1. How can I use this?

I can assess how my next-generation probiotic strains influence immune activation and tumor behavior during treatment using organoid technology. I can determine if these bacteria increase medication sensitivity, boost immune responses, or decrease cancer cell viability by subjecting tumor organoids to my NGPs or their metabolites in combination with chemotherapeutic treatments. I can also assess safety using organoids by seeing how they react to normal tissue organoids.

2. Why must I use this?

Mainly because organoid technology is more accessible and ethically controllable than animal models, and provides a degree of physiological realism that 2D cell cultures cannot.

3. When will I use this?

Organoids will be utilized in the mid-stage validation of my project, following the initial bacterial screening but prior to animal testing. To find out if my NGPs do improve immune activation and chemotherapeutic efficacy, I currently need a realistic model. When in vivo models are too expensive or too early, but simple cell culture is too limiting, organoids would be the best option.

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Currently, I'm working on the potential use of next-generation probiotics as chemotherapy adjuvants. Using high-throughput screening (HTS), I can easily test hundreds of probiotic strains or their metabolites to see which ones can boost immune responses or increase the efficacy of chemotherapy.

1. How can I use this technology?

- I can use HTS to quickly test thousands of probiotic strains or bacterial metabolites to find which ones activate immune cells, enhance chemotherapy responses, or reduce side effects. Instead of testing bacteria one by one, HTS helps me rapidly identify which probiotic candidates have the strongest and safest immune-modulating effects.

2. Why must I use this?

HTS guarantees that I don't overlook crucial strains or chemicals, enabling me to find the best probiotic options without having to guess. Other approaches are complex and tedious. On the other hand, HTS provides dependable, high-quality data quickly, which could help me identify the most promising NGP-based adjuvants for further testing.

3. When will I use this?

I will use this technology when I want to screen a variety of bacterial strains or their metabolites to determine which ones enhance immune function or improve chemotherapy results. Once I've determined which options are the best, I can test them further in subsequent studies.