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Essays on Genetic Engineering

What makes a good genetic engineering essay topic.

When it comes to writing a captivating genetic engineering essay, the topic you choose is paramount. It not only grabs the reader's attention but also allows for effective exploration of the subject matter. So, how can you brainstorm and select a standout essay topic? Here are some recommendations:

• Brainstorm: Kickstart your ideas by brainstorming topics related to genetic engineering. Consider the latest advancements, ethical concerns, controversial issues, or potential future applications. Jot down any ideas that come to mind.
• Research: Once you have a list of potential topics, conduct thorough research to gather relevant information and understand different perspectives. This will help you evaluate the feasibility and depth of each topic.
• Consider Interest: Choose a topic that genuinely piques your interest. Writing about something you are passionate about will make the entire process more enjoyable and motivate you to delve deeper into the subject matter.
• Relevance: Ensure that the chosen topic is relevant to genetic engineering. It should align with the scope of the subject and allow you to explore various aspects related to it.
• Uniqueness: Strive for a unique and imaginative topic that stands out from the ordinary. Steer clear of generic subjects and instead focus on specific areas or emerging trends within genetic engineering.
• Controversy: Controversial topics often generate more interest and discussion. Consider exploring ethical dilemmas, potential risks, or societal impacts of genetic engineering to add a thought-provoking element to your essay.
• Depth and Scope: Assess the depth and scope of each topic. Make sure it provides enough material for a comprehensive essay without being too broad or too narrow.
• Audience Appeal: Keep your target audience in mind. Choose a topic that would captivate readers, whether they are experts in the field or individuals with limited knowledge about genetic engineering.
• Originality: Strive for originality in your topic selection. Look for unique angles, lesser-known areas, or innovative applications of genetic engineering that can make your essay stand out.
• Personal Connection: If possible, choose a topic that connects with your personal experiences or future aspirations. This will enhance your engagement and make your essay more meaningful.

Igniting Thought: The Finest Genetic Engineering Essay Topics

Below are some of the most captivating genetic engineering essay topics to consider:

• Genetic Engineering and the Future of Human Evolution
• The Ethical Dilemmas of Designer Babies
• Genetic Engineering in Agriculture: Balancing Benefits and Concerns
• CRISPR-Cas9: Unleashing Revolutionary Potential in Genetic Engineering
• The Potential of Genetic Engineering in Cancer Treatment
• Genetic Engineering's Role in Creating Sustainable Food Sources
• Genetic Engineering and Animal Welfare: Navigating Ethical Considerations
• Genetic Engineering and its Impact on Biodiversity
• The Social and Economic Implications of Genetic Engineering
• Genetic Engineering's Influence on Human Longevity
• Enhancing Athletic Performance: The Power of Genetic Engineering
• Genetic Engineering Techniques for Disease Prevention and Treatment
• Genetic Engineering's Role in Environmental Conservation
• Genetic Engineering and the Preservation of Endangered Species
• The Psychological and Societal Effects of Genetic Engineering
• The Pros and Cons of Genetic Engineering for Non-Medical Purposes
• Exploring the Potential Risks and Benefits of Genetic Engineering in Space Exploration
• Genetic Engineering and the Creation of Biofuels
• The Morality of Genetic Engineering: Insights from Religious and Philosophical Perspectives
• Genetic Engineering's Role in Combating Climate Change

Thought-Provoking Genetic Engineering Essay Questions

Consider these stimulating questions for your genetic engineering essay:

• How does genetic engineering impact the concept of natural selection?
• What are the potential consequences of genetic engineering on human genetic diversity?
• Is it ethically justifiable to use genetic engineering for cosmetic purposes?
• How does genetic engineering contribute to the development of personalized medicine?
• What are the social implications of genetically modifying animals for human consumption?
• How does the use of genetic engineering in agriculture affect food security?
• Should genetic engineering be used to resurrect extinct species?
• What are the potential risks and benefits of genetically modifying viruses for medical purposes?
• How does genetic engineering influence the balance between individual rights and societal well-being?
• Can genetic engineering be the solution to eradicating genetic diseases?

Provocative Genetic Engineering Essay Prompts

Here are some imaginative and engaging prompts for your genetic engineering essay:

• Imagine a world where genetic engineering has eliminated all hereditary diseases. Discuss the potential benefits and drawbacks of such a scenario.
• You have been granted the ability to genetically engineer one aspect of yourself. What would you choose and why?
• Write a fictional story set in a future where genetic engineering is widespread and explore the consequences it has on society.
• Reflect on the ethical considerations of genetically modifying animals for entertainment purposes, such as creating glow-in-the-dark pets.
• Create a persuasive argument for or against the use of genetic engineering in enhancing human intelligence.

Answering Your Genetic Engineering Essay Queries

Q: Can I write about the history of genetic engineering?

A: Absolutely! Exploring the historical context of genetic engineering can provide valuable insights and set the foundation for your essay.

Q: How can I make my genetic engineering essay engaging for readers with limited scientific knowledge?

A: Simplify complex concepts and terminologies, provide relevant examples, and use relatable analogies to help readers grasp the information more easily.

Q: Can I express my personal opinion in a genetic engineering essay?

A: Yes, expressing your personal opinion is encouraged as long as you support it with logical reasoning and evidence from reputable sources.

Q: Are there any potential risks associated with genetic engineering that I should discuss in my essay?

A: Yes, incorporating a discussion on the potential risks and ethical concerns surrounding genetic engineering is essential to provide a balanced perspective.

Q: Can I include interviews or case studies in my genetic engineering essay?

A: Absolutely! Interviews or case studies can add depth and real-life examples to support your arguments and make your essay more compelling.

Remember, when writing your genetic engineering essay, let your creativity shine through while maintaining a formal and engaging tone.

The Ethics of Genetic Engineering in Human Enhancement

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Genetic Engineering: an Overview of The Dna/rna and The Crispr/cas9 Technology

Review of human germline engineering, positional cloning of genetic disorders, engineering american society: the lesson of eugenics, bioethical issues related to genetic engineering, cloning and ethical controversies related to it, genetic editing as a possibility of same-sex parents to have children, adhering to natural processes retains the integrity of a natural human race  , genetically modified organisms: soybeans, gene silencing to produce milk with reduced blg proteins, the role of crispr-cas9 gene drive in mosquitoes, the life of gregor mendel and his contributions to science, eugenics, its history and modern development, morphological operation hsv color space tree detetction, cytogenetics: analysis of comparative genomic hybridization and its implications, genetically engineered eucalyptus tree and crispr, review of the process of dna extraction, review of the features of the process of cloning, heterologous gene expression as an approach for fungal secondary metabolite discovery, review of the genetic algorithm searches.

Genetic engineering (also called genetic modification) is a process that uses laboratory-based technologies to alter the DNA makeup of an organism.

Genetic engineering as the direct manipulation of DNA by humans outside breeding and mutations has only existed since the 1970s. In 1972, Paul Berg created the first recombinant DNA molecules by combining DNA from the monkey virus SV40 with that of the lambda virus. The first field trials of genetically engineered plants occurred in France and the US in 1986, tobacco plants were engineered to be resistant to herbicides.

It is a set of technologies used to change the genetic makeup of cells, including the transfer of genes within and across species boundaries to produce improved or novel organisms. New DNA is obtained by either isolating and copying the genetic material of interest using recombinant DNA methods or by artificially synthesising the DNA. Used in research and industry, genetic engineering has been applied to the production of cancer therapies, brewing yeasts, genetically modified plants and livestock, and more.

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Genetic engineering articles within Nature Genetics

Article | 03 July 2023

Potent and uniform fetal hemoglobin induction via base editing

A comparison of fetal hemoglobin gene editing strategies using human sickle cell disease donor cells and in vivo transplantation finds that adenine base editing of the –175A>G site in the γ-globin gene promoters results in durable and potent expression.

• Thiyagaraj Mayuranathan
• , Gregory A. Newby
•  &  Jonathan S. Yen

Article 06 April 2023 | Open Access

Precise modulation of transcription factor levels identifies features underlying dosage sensitivity

SOX9 titration in neural crest cells identifies regulatory elements and genes with sensitive or buffered responses. Sensitive genes are enriched for craniofacial disorder genes phenocopying SOX9, suggesting differential sensitivity contributes to phenotypic specificity.

• Sahin Naqvi
• , Seungsoo Kim
•  &  Joanna Wysocka

Article | 23 March 2023

Thymidine nucleotide metabolism controls human telomere length

Genome-wide CRISPR screening identifies thymidine nucleotide metabolism as a key regulator of human telomere length. Thymidine supplementation promotes telomere elongation in cells derived from patients with telomere biology disorders.

• William Mannherz
•  &  Suneet Agarwal

Research Highlight | 13 March 2023

Engineering epigenetic inheritance

• Tiago Faial

Genome-scale characterization of transcription factors

• Michael Fletcher

Comment | 11 July 2022

30 years of progress from positional cloning to precision genome editing

Thirty years ago, I had the privilege of launching Nature Genetics , the first spin-off journal bearing the famous Nature logo. Spurred on by the Human Genome Project, there were high hopes for the new journal and indeed the future of human genetics. But there was little expectation that we would launch a science publishing franchise of more than 30 sister journals — or be able to therapeutically rewrite the faulty genomes of patients. Here, I reflect on the humble beginnings of Nature Genetics and 30 years of progress in genetics.

• Kevin Davies

Research Highlight | 07 September 2021

RNA demethylation for increased crop yields

Research Highlight | 06 August 2021

Sickle-cell anemia gene therapy

Hybrid potato genetics.

Article | 02 August 2021

Activation of γ-globin gene expression by GATA1 and NF-Y in hereditary persistence of fetal hemoglobin

Introduction of hereditary persistence of fetal hemoglobin variants into the γ-globin promoter by using CRISPR mutagenesis and editing provides insights into transcription factor interplay, with implications for gene therapies targeting this element.

• Phillip A. Doerfler
• , Ruopeng Feng
•  &  Mitchell J. Weiss

Perspective | 06 May 2021

Engineering three-dimensional genome folding

Recent technologies allow experimental manipulation of chromatin conformation. This Perspective discusses the insights obtained from gain-of-function studies that engineer the three-dimensional genome.

• , Jessica Lam
•  &  Gerd A. Blobel

Comment | 08 April 2021

A resource of targeted mutant mouse lines for 5,061 genes

The International Mouse Phenotyping Consortium reports the generation of new mouse mutant strains for more than 5,000 genes, including 2,850 novel null, 2,987 novel conditional-ready and 4,433 novel reporter alleles.

• Marie-Christine Birling
• , Atsushi Yoshiki
•  &  Stephen A. Murray

News & Views | 30 September 2020

Setting new boundaries with transcription and CTCF

How do boundary elements divide chromosomes into domains? A new study uses random genomic insertions to show how small genomic fragments can shape chromatin folding through the interplay of loop extrusion and compartmentalization. Spoiler: context matters.

• Erika C. Anderson
•  &  Elphège P. Nora

Article | 31 August 2020

Alteration of genome folding via contact domain boundary insertion

Insertion of a tissue-invariant chromatin domain boundary into 16 ectopic loci leads to various structural phenotypes, which depend on local chromatin features, CTCF binding and transcriptional status.

• , Peng Huang

Article | 01 June 2020

Selective Mediator dependence of cell-type-specifying transcription

Analysis with alleles encoding pharmacologically degradable Mediator subunits shows that Mediator acts as a global coactivator that facilitates transcription globally but is acutely required for cell-type-specific gene regulatory circuits.

• Martin G. Jaeger
• , Björn Schwalb
•  &  Georg E. Winter

Editorial | 28 February 2019

Brave new dialogue

The development of CRISPR–Cas technology and its applications in biomedical research have generated much excitement. If fully realized, this technology has the potential to help treat or prevent severe diseases. However, these tools also carry considerable risk if improperly used. The scientific community must promote constructive dialogue among its members and within society at large to ensure that research on genome editing is conducted responsibly.

Article | 27 July 2018

CRISPR–Cas9 genome editing in human cells occurs via the Fanconi anemia pathway

A coupled knockdown-editing screen shows that CRISPR–Cas9 editing in human cells requires the Fanconi anemia pathway, which acts by diverting double-strand break repair away from non-homologous end joining toward single-strand template repair.

• Chris D. Richardson
• , Katelynn R. Kazane
•  &  Jacob E. Corn

Editorial | 27 January 2016

Where genome editing is needed

The journal endorses the principle of transparency in the production of genome-edited crops and livestock as a precondition for the registration of a breed or cultivar, with no further need for regulation or distinction of these goods from the products of traditional breeding.

Research Highlights | 29 January 2014

A CRISPR method for genome-wide screening

• Brooke LaFlamme

Synthetic modeling of developmental enhancers

• Emily Niemitz

Research Highlights | 27 December 2013

ELABELA, a peptide hormone for heart development

Technical Report | 20 March 2011

Large-scale analysis of the regulatory architecture of the mouse genome with a transposon-associated sensor

Francois Spitz and colleagues report GROMIT, a Sleeping Beauty transposon–based system for mapping genetic regulatory architecture in mouse. GROMIT is a regulatory sensor that responds to the activity of nearby enhancers.

• , Orsolya Symmons
•  &  François Spitz

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January 16, 2019

Human Gene Editing: Great Power, Great Responsibility

Modifying the human germline has profound implications and must be approached with extraordinary care

By E. Paul Zehr

This article was published in Scientific American’s former blog network and reflects the views of the author, not necessarily those of Scientific American

We are at the point where our technology will soon surpass our humanity. It used to be that what we had in our jeans was just what we had in our genes. But we no longer are reliant on choosing our parents wisely. It was always going to happen. The new gene editing techniques were always going to be used to alter the genome in non-medically indicated cases. But it wasn’t anticipated we’d so soon have nontherapeutic application in human embryos.

On November 28, 2018, He Jiankui, from the Southern University of Science and Technology in Guangdong China, revealed that he had performed ex vivo gene editing on two human embryos. This was presented at the International Summit on Human Genome Editing in Hong Kong. It was not a therapeutic, medically indicated procedure, but, regardless, it was unethical and illegal in most countries.

 As an actual practicing scientist and as a human, I strongly advocate for advancement of science and leveraging our advances to enhance our species. Despite that, and somewhat ironically, when I began writing my most recent book, Chasing Captain America: How Advances in Science, Engineering, and Biotechnology Will Produce a Superhuman —a book explicitly focused on examining the science of altering human biology—I was skeptical about enhancing humanity. I challenged my perspective while writing and came to think we have an obligation to modify human form and function so we have the best chance to flourish on Earth and in space. Given the recently revealed experiments in which human embryos underwent nontherapeutic gene edits and were brought to term, we need to consider deeply the implications of this and ensure that what we do and how we proceed are grounded in ethical principles agreed upon by all of us.

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The idea of genetic engineering contained in gene editing is really no different in outcome than the pioneering work of Gregor Mendel in the mid-19th century and his detailed experiments with plants, particularly beans and peas. Mendel’s detailed observations of more than 10,000 plants taken over just about 10 years were published in 1866 and revealed the targeted changes in a living organism that could be obtained by breeding for desired characteristics.

Instead of producing desired characteristics, most of the biomedical work on gene therapy in our modern age focuses on therapeutic, medically indicated applications in inherited diseases and cancers. Many of these medical conditions arise because of dysfunctions in cellular metabolism, growth and viability. Of course, it is probably natural that along with the therapeutic application, there’s been interest in applications not aimed at “curing” disease but rather altering human performance in the otherwise “healthy.”

Gene editing techniques generally involve proteins that cut DNA, such as those employed in CRISPR-Cas9, transcription activator-like effector nucleases (TALENs) and zinc-finger nucleases. The most commonly used Cas enzyme, Cas9, comes from Streptococcus pyogenes —the one that gives you strep throat and was proven viable in mouse and human cells in 2013. The basic process is that the CRISPR molecule is programmed to search for a specific nucleotide sequence among the 3 billion in the human genome. Once the correct sequence is identified, CRISPR unwinds the coils of DNA coils and “snips” the sequence out of the strand. DNA strands are then repaired in the case of a gene deletion, or, for an insertion, a new sequence can be included to alter the genome.

Performed in an embryonic germ line cell, an egg or a sperm cell, gene “edits” will be part of the genetic code that goes to the next generation. But there can be errors—in other words, editing more than intended—with targeting associated with the guide RNA used to target the deletions. It is the presence of these “off-target repeats” that indicates extreme caution and a need for better regulation before techniques like CRISPR can have safe clinical application.

As such, we as scientists and society must also balance the potential good associated with new techniques and the prospect of doing something just because we could. Gene editing places great power over altering the fundamental principles of biology, and our whole society needs to part of the discussion on what is okay to do and what is not. And we need to move quickly but not in a hurry.

It’s critical to think about the path ahead—which one to take and to where—before we arrive. Scientists and engineers at work right now are working to enable the realization of our common futures. But guiding the implementation of that future is the right and responsibility of us all and cannot be entrusted exclusively to those at work in the field and laboratories, nor to those who attempt to regulate their work, our lawmakers and bureaucrats.

The future we invent can be bright—but there are strings attached. The most important string is that we need input from as many sectors in our society as possible. The decisions that are made will literally affect the future of our species and cannot be made in isolation from our society as a whole.

Science works as a machine of chance effects with experimental outcomes; tested against a backdrop of random occurrences and biological evolution is the emergence of chance survival characteristics expanding over millions of years. There is a pace and timing to adaptations. Yet, any modifying of the human germ line—editing sperm or egg cells—has direct implications for the next generation and must be done carefully in light of regulations specifically addressing this kind of experimentation. In many countries there is a de facto moratorium on human germ line and embryo editing because such work is illegal. It is also completely unethical, not least of all because of lack of consent.

Eike-Henner Kluge from the University of Victoria has written that “germ line alteration would be performed without the consent of those who are most affected: namely, future generations.” And C.S. Lewis, when he wasn’t enthralling us with the Chronicles of Narnia, wrote in 1965’s The Abolition of Man that if a society gains power to make descendants “what it pleases, all men who live after it are patients of that power … the rule of a few hundreds of men over billions upon billions of men.”

All of us citizens, scientists, engineers and future users of human enhancement methodologies must proceed with conviction but also caution, with purpose but also extreme care. It’s critical to appreciate the implications of the power of science as articulated by Richard Dawkins that “science is the most powerful way to do whatever it is you want to do. If you want to do good, it’s the most powerful way of doing good. If you want to do evil, it’s the most powerful way to do evil.” Never before have we—or any other species on this planet—had such influence and so much power over the fundamental nature of our own biology.

The nontherapeutic use of gene editing on human embryos was and remains unethical and illegal on every level. Yet, now we need to leverage attention on gene editing and human enhancement into a real conversation about the future our species. As the late Stan Lee wrote back in 1962 in Amazing Fantasy , the first comic book featuring Spider-Man, “with great power there must also come—great responsibility!”

Both must be exercised judiciously here and now in real life.

Arguing For and Against Genetic Engineering

Harvard philosopher Michael Sandel recently spoke at Stanford on the subject of his new book, The Case against Perfection: Ethics in the Age of Genetic Engineering. He focused on the “ethical problems of using biomedical technologies to determine and choose from the genetic material of human embryos,” an issue that has inspired much debate.

Having followed Sandel’s writings on genetic enhancement for several years, I think that this issue deserves special thought. For many years, the specter of human genetic engineering has haunted conservatives and liberals alike. Generally, their main criticisms run thus:

First, genetic engineering limits children’s autonomy to shape their own destinies. Writer Dinesh D’Souza articulates this position in a 2001 National Review Online article: “If parents are able to remake a child’s genetic makeup, they are in a sense writing the genetic instructions that shape his entire life. If my parents give me blue eyes instead of brown eyes, if they make me tall instead of medium height, if they choose a passive over an aggressive personality, their choices will have a direct, lifelong effect on me.” In other words, genetic enhancement is immoral because it artificially molds people’s lives, often pointing their destinies in directions that they themselves would not freely choose. Therefore, it represents a fundamental violation of their rights as human beings.

Second, some fear that genetic engineering will lead to eugenics. In a 2006 column, writer Charles Colson laments: “British medical researchers recently announced plans to use cutting-edge science to eliminate a condition my family is familiar with: autism. Actually, they are not ‘curing’ autism or even making life better for autistic people. Their plan is to eliminate autism by eliminating autistic people. There is no in utero test for autism as there is for Down syndrome…[Prenatal] testing, combined with abortion-on-demand, has made people with Down syndrome an endangered population…This utilitarian view of life inevitably leads us exactly where the Nazis were creating a master race. Can’t we see it?” The logic behind this argument is that human genetic enhancement perpetuates discrimination against the disabled and the “genetically unfit,” and that this sort of discrimination is similar to the sort that inspired the eugenics of the Third Reich.

A third argument is that genetic engineering will lead to vast social inequalities. This idea is expressed in the 1997 cult film Gattaca, which portrays a society where the rich enjoy genetic enhancements—perfect eyesight, improved height, higher intelligence—that the poor cannot afford. Therefore, the main character Vincent, a man from a poor background who aspires to be an astronaut, finds it difficult to achieve his goal because he is short-sighted and has a “weak heart.” This discrepancy is exacerbated by the fact that his brother, who is genetically-engineered, enjoys perfect health and is better able to achieve his dreams. To many, Gattaca is a dystopia where vast gaps between the haves and have-nots will become intolerable, due to the existence of not just material, but also genetic inequalities.

The critics are right that a world with genetic engineering will contain inequalities. On the other hand, it is arguable that a world without genetic engineering, like this one, is even more unequal. In Gattaca, a genetically “fit” majority of people can aspire to be astronauts, but an unfortunate “unfit” minority cannot. In the real world, the situation is the other way round: the majority of people don’t have the genes to become astronauts, and only a small minority with perfect eyesight and perfect physical fitness—the Neil Armstrong types—would qualify.

The only difference is that in the real world, we try to be polite about the unpleasant realities of life by insisting that the Average Joe has, at least theoretically, a Rocky-esque chance of becoming an astronaut. In that sense, our covert discrimination is much more polite than the overt discrimination of the Gattaca variety. But it seems that our world, where genetic privilege exists naturally among a tiny minority, could conceivably be less equal (and less socially mobile) than a world with genetic engineering, where genetic enhancements would be potentially available to the majority of people, giving them a chance to create better futures for themselves. Supporters of human genetic engineering thus ask the fair question: Are natural genetic inequalities, doled out randomly and sometimes unfairly by nature, more just than engineered ones, which might be earned through good old fashioned American values like hard work, determination, and effort?

“But,” the critics ask, “wouldn’t genetic engineering lead us to eugenics?” The pro-genetic engineering crowd thinks not. They suggest that genetic engineering, if done on a purely decentralized basis by free individuals and couples, will not involve any form of coercion. Unlike the Nazi eugenics program of the 1930s, which involved the forced, widespread killing of “unfit” peoples and disabled babies, the de facto effect of genetic engineering is to cure disabilities, not kill the disabled. This is a key moral difference. As pointed out by biologist Robert Sinsheimer, genetic engineering would “permit in principle the conversion of all the ‘unfit’ to the highest genetic level.” Too often, women choose to abort babies because pre-natal testing shows that they have Down syndrome or some other ailment. If anything, genetic engineering should be welcomed by pro-life groups because by converting otherwise-disabled babies into normal, healthy ones, it would reduce the number of abortions.

In addition, the world of Gattaca, for all its faults, features a world that, far from being defined along Hitler-esque racial lines, has in fact transcended racism. Being blond-haired and blue-eyed loses its racially elitist undertones because such traits are easily available on the genetic supermarket. Hair color, skin color, and eye color become a subjective matter of choice, no more significant than the color of one’s clothes. If anything, genetic engineering will probably encourage, not discourage, racial harmony and diversity.

It is true that genetic engineering may limit children’s autonomy to shape their own destinies. But it is equally true that all people’s destinies are already limited by their natural genetic makeup, a makeup that they are born with and cannot change. A short person, for example, would be unlikely to join the basketball team because his height makes it difficult for him to compete with his tall peers. An ugly person would be unable to achieve her dream of becoming a famous actress because the lead roles are reserved for the beautiful. A myopic kid who wears glasses will find it difficult to become a pilot. A student with an IQ of 75 will be unlikely to get into Harvard however hard he tries. In some way or another, our destinies are limited by the genes we are born with.

In this sense, it is arguable that genetic engineering might help to level the playing field. Genetic engineering could give people greater innate capacity to fulfill their dreams and pursue their own happiness. Rather than allow peoples’ choices to be limited by their genetic makeup, why not give each person the capability of becoming whatever he or she wants to, and let his or her eventual success be determined by effort, willpower, and perseverance? America has long represented the idea that people can shape their own destinies. To paraphrase Dr. King, why not have a society where people are judged not by the genes they inherit, but by the content of their character?

Looking at both sides, the genetic engineering controversy does raise questions that should be answered, not shouted down. Like all major scientific advances, it probably has some negative effects, and steps must be taken to ameliorate these outcomes. For example, measures should also be taken to ensure that genetic engineering’s benefits are, at least to some extent, available to the poor. As ethicists Maxwell Mehlman and Jeffrey Botkin suggest in their book Access to the Genome: The Challenge to Equality, the rich could be taxed on genetic enhancements, and the revenue from these taxes could be used to help pay for the genetic enhancement of the poor. To some extent, this will help to ameliorate the unequal effects of genetic engineering, allowing its benefits to be more equitably distributed. In addition, caution must be taken in other areas, such as ensuring that the sanctity of human life is respected at all times. In this respect, pro-life groups like Focus on the Family can take a leading role in ensuring that scientific advances do not come at the expense of moral ethics.

At the same time, we should not allow our fear of change to prevent our society from exploring this promising new field of science, one that promises so many medical and social benefits. A strategy that defines itself against the core idea of scientific progress cannot succeed. Instead of attempting to bury our heads in the sand, we should seek to harness genetic engineering for its positive benefits, even as we take careful steps to ameliorate its potential downsides.

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Genetic Engineering by DNA Recombineering

Louis j. papa, iii.

1 Massachusetts Institute of Technology, Department of Chemistry, 77 Massachusetts Avenue, Cambridge, Massachusetts 02139

Matthew D. Shoulders

Recombineering inserts PCR products into DNA using homologous recombination. A pair of short homology arms (50 base pairs) on the ends of a PCR cassette target the cassette to its intended location. These homology arms can be easily introduced as 5′ primer overhangs during the PCR reaction. The flexibility to choose almost any pair of homology arms enables the precise modification of virtually any DNA for the purposes of sequence deletion, replacement, insertion, or point mutation. Recombineering often offers significant advantages relative to previous homologous recombination methods that require the construction of cassettes with large homology arms and relative to traditional cloning methods that become intractable for large plasmids or DNA sequences. However, the tremendous number of variables, options, and pitfalls that can be encountered when designing and performing a recombineering protocol for the first time introduce barriers that can make recombineering a challenging technique for new users to adopt. This article focuses on three recombineering protocols we have found to be particularly robust, providing a detailed guide for choosing the simplest recombineering method for a given application, and for performing and troubleshooting experiments.

INTRODUCTION

Recombineering is a versatile genetic engineering method that can be used to introduce deletions, insertions, gene replacements or point mutations virtually anywhere in a DNA sequence and is particularly useful for editing large Bacterial Artificial Chromosomes (BAC), DNA virus genomes, or bacterial genomes. The method utilizes the temporary expression of lambda phage genes ( red αβγ ) that facilitate recombination between the target BAC or target genome and a DNA targeting cassette that is introduced into the cell through electroporation. Because only about 50 base pairs (bp) of homology are required to facilitate recombination, the homology arms of a recombineering targeting cassette can be conveniently introduced as primer overhangs in a single PCR step ( Murphy, 2016 ). Other homologous recombination-based methods generally require 500–1000 bp homology arms that need to be appended to a targeting cassette through time-consuming restriction cloning or difficult overlap extension PCR ( Hamilton, Aldea, Washburn, Babitzke, & Kushner, 1989 ; Kong, Yang, & Geller, 1999 ; Uil et al., 2011 ; Winans, Elledge, Krueger, & Walker, 1985 ). Another major advantage of recombineering is the ability to modify plasmids of virtually any size. Other popular in vitro DNA editing methods, such as endonuclease cloning or in vitro assembly-based techniques rely on transferring the recombinant DNA plasmid into E. coli after the modifications have been made in a process known as “transformation”. The efficiency of transformation and the ability to modify DNA plasmids in vitro decreases dramatically for plasmids greater than 10 kb ( Inoue, Nojima, & Okayama, 1990 ; Siguret, Ribba, Chérel, Meyer, & Piétu, 1994 ), rendering most in vitro methods impractical for modifying large plasmids. Because recombineering modifies plasmids that already reside in the E. coli cell, it circumvents the transformation bottleneck. Additionally, recombineering creates modifications without the need to cut or ligate DNA, thereby avoiding other challenges associated with in vitro methods when modifying very large plasmids, such as the lack of unique restriction sites or the difficulty of amplifying large DNA fragments by PCR ( Cheng, Fockler, Barnes, & Higuchi, 1994 ).

Typical applications of recombineering include modifying the E. coli genome itself or the genomes of large human or animal viruses as BACs in E. coli that are later reintroduced into the host organism ( Berman et al., 2018 ; Moore et al., 2018 ; Narayanan & Chen, 2011 ). The modified genomes can then produce genetically-engineered virions in a process known as “viral rescue”. Recombineering is also ideal for editing BACs with large fragments of human or animal genomes, such that these modified fragments can then be used as targeting cassettes to modify human or animal cells via non-recombineering homologous recombination ( Murphy, 2016 ). Note that, in human and animal cells, long homology arms are required and thus targeting cassettes must necessarily be large ( Baker et al., 2017 ; Hasty, Rivera-Pérez, & Bradley, 1991 ). Importantly, recombineering is not limited to modifying the E. coli genome, viral genomes, or large targeting cassettes. Rather, it can be used whenever a large DNA construct needs to be modified in E. coli .

There are many decisions to make when choosing an application-appropriate recombineering strategy. In particular, the multitude of available recombineering-based methods and their many variations can make it daunting for a new user to choose the best option for a specific purpose. This article aims to distill the many possible iterations down to just three simple protocols that, based on our extensive experience engineering adenovirus vectors ( Berman et al., 2018 ; Wong et al., 2018 ), E. coli genomes ( Moore et al., 2018 ), and other DNA sequences, function reliably and efficiently. Further, although recombineering can be incredibly simple and efficient to apply, it is also sensitive to many critical experimental parameters. Thus, each protocol provides, as a companion resource, detailed annotations for avoiding common pitfalls. In order of increasing difficulty and required time, the protocols are: Basic Protocol 1 —One-Step Recombineering ( Datsenko & Wanner, 2000 ; Yu et al., 2000 ), Basic Protocol 2—Direct and Inverted Repeat stimulated Excision (DIRex) Recombineering ( Näsvall, 2017 ), and Basic Protocol 3 —Standard Two-Step Recombineering ( H. Wang et al., 2014 ). As explained below, the ideal choice of protocol depends on factors such as the type of modification to be made, the size of the modification to be made, and the number of modifications to be made on a given BAC or genome. Mastery of the protocols and concepts described here should provide a strong foundation for understanding, utilizing, and adapting these and other recombineering-based methods.

STRATEGIC PLANNING

Under standard double-stranded DNA recombineering conditions, only 1 in 10,000–100,000 cells yields successful recombinants that incorporate the targeting cassette at the desired location ( Datta, Costantino, Zhou, & Court, 2008 ). This frequency is far too low to reasonably screen colonies one-by-one for recombinants, so one must employ selection-based methods where only the rare recombinants survive after selection. Unfortunately, most modifications do not significantly impact the survival of the E. coli cell and therefore cannot be selected for directly. Thus, either an antibiotic resistance gene must be included in the modification to yield a “marked” modification ( Basic Protocol 1 ), or a two-step selection-counterselection method must be employed to yield a “seamless” modification in which the final recombinant does not contain an antibiotic resistance gene ( Basic Protocol 2 or 3 ). To choose the simplest protocol for a desired modification, refer to the decision tree in Figure 1A . Detailed potential uses and time requirements for each of the protocols are provided in Figure 1B .

Protocol selection guide. ( A ) Decision tree for choosing the quickest, simplest protocol for making a desired type of modification. ( B ) The potential uses and time requirements of each protocol.

For all three Basic Protocols described in this article, it is essential to obtain the target DNA in a BAC unless the E. coli genome itself is to be modified. BACs have low-copy origins (1–2 copies per cell), which is necessary ensure that any plasmids that are modified by recombineering will be isolated in a given cell without any copies of the parent plasmid. If a high-copy plasmid were used as the target, successful recombinants would be mixed with unmodified plasmid within a single cell, even after selection pressure is applied ( Thomason, Costantino, Shaw, & Court, 2007 ). For Basic Protocol 1 , the mixture would need to be re-transformed to purify the successful recombinant. For Basic Protocols 2 and 3 , the counterselection step would be impossible. Fortunately, it is straightforward to replace any plasmid origin with the BAC origin using one-step recombineering (via Basic Protocol 1 ). We note that a significant drawback of BACs is that their low copy-number makes it very difficult to purify large quantities of DNA, which is often necessary for downstream applications such as DNA transfection into mammalian cells. Therefore, once all of the desired modifications have been made, it is often useful to replace the low-copy BAC origin with a high-copy pUC origin using one-step recombineering (via Basic Protocol 1 ). We further note that, while there are some BACs that can switch to high-copy numbers upon arabinose induction by utilizing a second origin, oriV ( Westenberg, Bamps, Soedling, Hope, & Dolphin, 2010 ; Wild, Hradecna, & Szybalski, 2002 ), all the protocols described here use arabinose induction for other purposes. If oriV -containing BACS were used, the consequence would be undesired high-copy replication of the BACs. Thus, it is essential to ensure that any target BAC employed does not contain oriV .

BASIC PROTOCOL 1

One-step recombineering for marked insertions, replacements or deletions.

Introducing a marked modification is the most straightforward way to create DNA insertions, deletions or replacements—large or small—and can be accomplished quickly in a single step ( Datsenko & Wanner, 2000 ). To introduce a marked modification, an antibiotic resistance gene needs to either replace the region that is to be deleted or be included with any regions that are to be inserted, such that recombinants can be selected for by streaking cells on an antibiotic-containing plate ( Fig. 2 ). Marked modifications can be useful for when only one or a few deletions, insertions, or replacements need to be introduced in a given piece of DNA. Nonetheless, the number of sequential modifications that can be introduced is necessarily limited by the requirement for a new antibiotic resistance gene at each step ( Thomason, Sawitzke, Li, Costantino, & Court, 2014 ). Finally, we note that although various scales can be used, we strongly recommend (and describe below) performing the protocol on a 1 mL scale for ease of sample processing and using heat blocks, as described in previous work ( H. Wang et al., 2014 ).

Schematic of one-step recombineering in which a kanamycin resistance gene ( kan R ) is used to select for recombinants and thereafter becomes part of the modification.

Target BAC (supplied by user)

psc101-gbaA plasmid (see Table 1 )

Plasmids used in these protocols

Targeting primers (designed and ordered by user)

Sequencing primers (designed and ordered by user)

R6K-kan-ccdB plasmid (see Table 1 )

Template plasmid (used for insertions; supplied by user)

pBeloBAC11loxP2272 (see Table 1 )

pcDNA-DEST40 (see Table 1 )

1.7 mL microcentrifuge tubes (e.g., VWR 87003–294)

Bunsen burner and gas line

18-gauge needles (e.g., Becton Dickinson 305195)

Sterile 10 μL pipette tips (e.g., VWR 89079–466)

Sterile 200 μL pipette tips (e.g., VWR 89079–460)

Sterile 1000 μL pipette tips (e.g., VWR 89079–472)

Lysogeny broth (LB) media (see recipe)

1000X streptomycin stock solution (see recipe)

DH10B E. coli cells (Invitrogen 18297010) or other target E. coli strain

50 mL conical tubes (e.g., Cellstar 227261)

0.1 cm electroporation cuvettes (e.g., Bio-Rad 1652083)

Kimwipes (Kimberly-Clark 34155)

Super optimal broth with catabolic repression (SOC) media (see recipe)

LB agar plates with 1X chloramphenicol and 1X tetracycline (see recipe)

9” soda-lime glass Pasteur pipettes (e.g., VWR 14672–380)

1000X tetracycline stock solution (see recipe)

1000X chloramphenicol stock solution (see recipe)

Sterile 50% glycerol (see recipe)

10% rhamnose stock solution (see recipe)

LB agar plates with 1X kanamycin (see recipe)

14 mL sterile culture tubes (e.g., VWR 60818–703)

10 mL serological pipette (e.g., Cellstar 607180)

1000X kanamycin stock solution (see recipe)

Miniprep DNA isolation kit (e.g., Omega Bio-tek Plasmid Mini Kit D6942–01)

Isopropanol (e.g., Macron 3032–16)

70% ethanol (e.g., Koptec V1401)

Restriction endonucleases (selected by user; e.g., New England Biolabs)

Agarose (e.g., Lonza 50004)

0.5X TBE buffer (see recipe)

Paper towels

GelGreen Nucleic Acid Gel Stain (Biotium 41005)

Quick-Load Purple 1 kb DNA Ladder (New England Biolabs N0552S)

6X Gel Loading Dye (New England Biolabs B7024S)

DNA Editing Software (e.g., Snapgene, ApE, Serial Cloner)

P10 pipettor (e.g., VWR 89079–962)

P200 pipettor (e.g., VWR 89079–970)

P1000 pipettor (e.g., VWR 89079–974)

Shaking heat block for 1.5 mL tubes (e.g., Eppendorf Thermomixer F1.5 5384000020)

UV-Vis spectrometer (e.g., Thermo Fisher Nanodrop 2000c)

Ice maker and ice buckets

Refrigerated microcentrifuge ≥16,000 × g (e.g., Thermo Fisher Heraeus Fresco 21 75002426)

Electroporator (Bio-Rad Micropulser 1652100)

30 °C incubator (e.g., VWR Gravity Convection Incubator 89511–422)

Pipette gun (e.g., Drummond Portable Pipet-Aid XP 4–000-101)

30 °C shaking incubator (e.g., Thermo Fisher MaxQ 4000 SHKE4000)

Precision balance (e.g., Mettler Toledo ME1002TE 30216559)

250 mL glass Erlenmeyer flask (e.g., VWR 10536–914)

Agarose gel electrophoresis system (e.g., Bio-Rad Mini-Sub Cell GT System 1664401)

Microwave oven

Mini-gel caster (e.g., Bio-Rad 1704422)

Blue LED transilluminator (e.g., Maestrogen LED Transilluminator SLB-01W)

Design targeting primers and create the targeting cassette

Primer design example for a marked insertion via one-step recombineering. ( A ) The left and right targeting primers amplify the region to be inserted, which contains a kanamycin resistance marker ( kan R ). The left and right targeting primers have overhangs that are homologous to the regions flanking the target region to be replaced. ( B ) The left homology arm (red underlined) is copied from the top strand while the right homology arm (blue underlined) is copied from the bottom strand. ( C ) The left primer that amplifies the modification from the template plasmid (brown underlined) is copied from the top strand, and the right primer that amplifies the modification from the template plasmid (green underlined) is copied form the bottom strand. ( D ) The homology arms are appended to the primers to form the full targeting primers. Note that any sequences copied from the bottom strand are reversed here to be represented in the 5′ to 3′ direction.

• Immediately to the right of the region to be deleted or replaced, highlight 50 bp, not including the region to be deleted or replaced, and copy or mark the sequence from the BOTTOM strand for the right homology arm. Ensure that the sequence copied and designated as the right homology arm is the bottom strand in the 5′ to 3′ direction (i.e., the reverse complement of the TOP strand). In the case of an insertion, the 3′ ends of the left homology arm and the right homology arm will be adjacent to each other if the homology arms are properly designed.

The protocols in this article use kanamycin resistance for selection of recombinants because it works reliably. Other antibiotic resistance genes can also be used for selection. However, ampicillin is not as reliable as kanamycin for killing susceptible bacteria because the β-lactamase enzyme that confers ampicillin resistance is secreted and destroys extracellular antibiotic, allowing some nearby susceptible cells to survive and grow ( Medaney, Dimitriu, Ellis, & Raymond, 2016 ). Therefore, we recommend against using ampicillin resistance for routine recombineering.

Primer sequences

• An insertion or replacement: Create or obtain a plasmid that has the genes or region to be inserted into the target. The region to be inserted must include an antibiotic resistance gene for selection. Design primers that have melting temperatures ( T m ) of ~60 °C—note that most DNA editing softwares display the T m as DNA is highlighted—and that amplify the entire region to be inserted ( Fig. 3C ) along with the antibiotic resistance gene ( Sawitzke et al., 2013 ). In the primer design, append the left homology arm to one primer and the right homology arm to the other primer ( Fig. 3D ). Note that the choice of which homology arm to append to which primer will determine the ultimate orientation of the insertion. Once the primers are obtained, use them in conjunction with the template plasmid to create the targeting cassette (see Support Protocol 1 ).

Replacing the origin of a high-copy plasmid with the BAC origin nearly always results in the formation of a heteromultimer in which multiple high-copy plasmids and BACs fuse together into one large plasmid with mixed origins. Any successful recombinant cells also likely contain unmodified parent plasmids. Heteromultimers and/or mixtures are unsuitable targets for further recombination. Therefore, it is generally necessary to “remonomerize” and isolate the recombinant BAC via restriction digestion as described in Support Protocol 4 . Alternatively, target DNA can be ligated to the origin of pBeloBac11loxP2272 using standard restriction endonuclease cloning.

• Converting a BAC into a high-copy pUC plasmid: Obtain and use primers pUC.For and pUC.Rev ( Table 2 ) in conjunction with pcDNA-DEST40 to create the targeting cassette (see Support Protocol 1 ). The pUC.For and pUC.Rev primers already have the necessary homology arms, assuming the BAC is derived from pBeloBac11loxP2272. If in doubt, double-check the vector sequence to ensure that the homology arms in pUC.For and pUC.Rev exist in the target BAC and flank the BAC origin. Note that this targeting cassette confers ampicillin resistance. The pcDNA-DEST40 plasmid is particularly useful not only because it provides a template for the pUC origin, but also because it encodes the toxic ccdB gene. If any template plasmid enters the cell during the recombineering electroporation step, the ccdB gene will kill that cell and prevent false positives resulting from template plasmid replication. Templates other than pcDNA-DEST40 can be used to generate a targeting cassette with the pUC origin, but then precautions should be taken to inactivate the template (see Support Protocol 1 ) and the user may need to redesign the pUC.For and pUC.Rev primers, depending on the alternate template chosen.

Replacing the origin of a BAC with the pUC origin almost always results in the formation of a multimers in which multiple pUCs and/or BACs fuse together into one large plasmid. Multimers may be unsuitable for later applications. Therefore, it is wise to “remonomerize” the recombinant pUC via restriction digestion as described in Support Protocol 4 .

In general, sequencing primers should be ~200 bp away from the homology arms so that any PCR products generated of the region will be at least 400 bp in length ( Murphy, 2016 ; Thomason et al., 2014 ). PCR products shorter than 400 bp can be difficult to visualize on and purify from a standard agarose gel. In addition to PCR amplifying the region of interest, the sequencing primers will also be used in the subsequent Sanger sequencing reaction. The first 50–100 bp of a Sanger sequencing trace tends to be noisy and unreliable for interpreting the sequence. Placing the sequencing primer 200 bp upstream of the homology arm ensures that the trace is reliable in the region of the recombination junction, thus allowing confirmation of successful recombination.

Transforming the target BAC and psc101-gbaA recombineering plasmid into DH10B E. coli

Heating the needle sterilizes it and helps melt a needle-sized hole into the plastic. This hole is critical for proper aeration and growth of the bacterial culture. Never hold the microcentrifuge tube in your hand while poking a hole in the lid. Instead, secure the tube in a rack.

• 6. If the target BAC and the psc101-gbaA plasmid are already both present in the same E. coli strain, proceed to step 23. If the intent is to engineer the E. coli genome instead of a target BAC, proceed to step 7 but omit the addition of chloramphenicol to all agar plates and media in subsequent steps.

When preparing or manipulating bacterial cultures in general, use sterile technique to prevent contamination. Keep tubes, bottles, and tips boxes closed and only open momentarily as needed. Everything that comes in contact with the culture must be sterile. Perform any manipulations that require sterility near the base of a Bunsen burner or inside a biosafety cabinet to minimize the chance of airborne contamination.

• 8. Inoculate the media with DH10B E. coli using either a stab from a glycerol stock or a colony from a plate streaked with DH10B E. coli . Place the tube in a shaking heat block set to 30 °C and 1000 r.p.m. and incubate overnight for ~16 hours.

After 2.5 hours, the culture should appear slightly cloudy with an OD 600 of about 0.6, but it is not necessary to measure the OD every time. The slightly cloudy culture is in “mid-log” phase, meaning the cells are growing rapidly and are displaying logarithmic growth. Mid-log phase is when the cells are most “competent,” meaning that the physiology of the cells is most amenable to DNA uptake through electroporation ( Calvin & Hanawalt, 1988 ). Do not try to electroporate cells with an OD of 1 or greater (a culture that appears opaque in a 1.7 mL microcentrifuge tube), as they will not efficiently uptake DNA. It is also not sufficient to dilute a “stationary” culture (i.e., one that has stopped growing) such as an overnight to an OD of 0.6 directly. It is not the actual concentration of cells that matters, but rather their state, which is dependent on their rate of growth at the time they are harvested.

• 10. Place the culture on ice to arrest growth. Next, pellet the cells at 10,000 × g in a refrigerated microcentrifuge at 4 °C for 1 minute. Carefully remove the media without disturbing the pellet.
• 11. Gently resuspend the pellet in 1 mL of ice-cold molecular biology-grade water by pipetting several times. Pellet again at 10,000 × g in a refrigerated microcentrifuge at 4 °C for 1 minute.

The purpose of these washing steps is to remove the salt and other electrolytes surrounding the cells, which is critical. During the later electroporation step, the cells will be placed into an electroporation cuvette, which essentially acts as a capacitor. In the cuvette, the cells will be briefly exposed to a strong electric field, which will induce the formation of temporary pores in the cell membrane through which DNA can enter. If there is too much salt or other electrolytes surrounding the cells, the sample will conduct electricity between the positive and negative plates of the cuvette during the electroporation, which will kill the cells.

Often there is ~20 μL of residual water around the pellet, in which case it is unnecessary to add additional water to the tube.

It is critical that the target BAC and psc101-gbaA plasmid were eluted from their purification columns in molecular biology-grade water or 0.2X elution buffer (see recipe). The salt concentration of the elution buffer in most plasmid purification kits will cause the sample to conduct electricity and kill the cells. If the DNA sample was eluted in 1X elution buffer or the salt concentration is unknown, the sample can easily be desalted using a PCR clean-up kit, such as the E.Z.N.A. Omega Bio-tek Cycle Pure Kit. Follow manufacturer instructions, but be sure to elute in molecular biology-grade water or 0.2X elution buffer for the final step. Also, do not add more than 5 μL of DNA sample to the cells, as doing so often causes the samples to conduct electricity even if the DNA samples were eluted in molecular biology-grade water or 0.2X elution buffer.

It is important to have everything set up as described before electroporating, because all the subsequent steps need to be completed very quickly.

Do not add more than 40 μL of cells to the cuvette. Adding large volumes decreases the resistance of the sample and can cause the sample to conduct electricity.

While it is important to move quickly, do not slam the electrocuvette into the electroporation chamber. Doing so often launches the sample out from between the plates of the electrocuvette, resulting in a failed electroporation because the cells were not exposed to the strong electric field between the plates.

The time between electroporation and resuspension should ideally not exceed 10 seconds. Fewer and fewer cells will survive the longer it takes to resuspend the cells ( Dower, Miller, & Ragsdale, 1988 ). While SOC media helps maximize the number of cells that survive, it is not strictly necessary and can be substituted with LB media.

If the electroporation was successful, the pulse length displayed on the screen should be between 4.80 and 6.00 ms. If the pulse length is less than 4.80 ms or if the screen says “Arc”, it means that the voltage applied to the electrocuvette decayed too quickly because significant current passed through the cells. In this case, repeat the electroporation with new electrocompetent cells and add a smaller quantity of DNA. It may be necessary to desalt the DNA again or wash the cells an extra time. It is often useful to prepare a few batches of electrocompetent cells in parallel in case the electroporation needs to be repeated a few times before it is successful.

• 19. Return the resuspended cells to the 1.7 mL microcentrifuge they came from and incubate them in the heat block at 30 °C for 1 hour at 1,000 r.p.m.

It is essential to provide the cells enough time (~1 hour) to recover and express any antibiotic resistance genes before spreading them on an antibiotic containing plate.

Melt the thin end of a 9” soda-lime Pasteur pipette into a hockey stick shape using the Bunsen burner. Do not use borosilicate Pasteur pipettes, as they do not melt easily. Give the hockey stick 10 seconds to cool, then use it to spread the cells evenly on the plate. Allow the plate to dry with the lid off near the base of the Bunsen burner. The updraft created by the Bunsen burner prevents dust or other contaminants from settling onto the plate. Cells can also be spread on the plate with sterile glass beads or a sterile 1000 μL plastic pipette tip, but best results are usually achieved with a glass hockey stick.

• 22. After the plate has dried, meaning there is no more running liquid on top of the agar, incubate it at 30 °C for ~18 hours or until colonies appear.

If colonies do not appear after 2 days, return to step 5. If the DNA samples are of sufficient quality (>20 ng/μL with an A260/280 of ~1.8–1.9), it should be possible to obtain cells that simultaneously take up the psc101-gbaA plasmid and the target BAC. However, if electroporating both plasmids at once is not successful, instead electroporate the target BAC alone into DH10B cells first, then repeat the process to electroporate the psc101-gbaA cells into DH10B cells that contain the target BAC. An alternative is to maintain a strain of DH10B cells that already contain the psc101-gbaA recombineering plasmid, in which case only the target BAC needs to be electroporated into the DH10B strain that already contains psc101-gbaA. When troubleshooting, electroporating 100 ng of any high-copy positive control plasmid less than 6000 base pairs in length (which should yield thousands of colonies) can help to evaluate whether the electroporation apparatus or electrocompetent cells are the source of any problems.

Recombineering with the targeting cassette

The replication of the psc101-gbaA plasmid is temperature-sensitive. Thus, it is important not to exceed 30 °C for any extended period of time to prevent loss of the psc101-gbaA plasmid.

• 24. Use 40 μL of the overnight culture, which should now appear opaque, to inoculate a new 1 mL LB culture with 1X chloramphenicol and 1X tetracycline. Incubate the new culture at 30 °C for exactly 2.0 hours. While the culture is growing, chill 50 mL of molecular biology-grade water in a 50 mL conical tube on ice. Optionally, thoroughly mix 500 μL of the overnight culture with 500 μL of sterile 50% glycerol in a sterile 1.7 mL microcentrifuge tube (without a hole in the lid) and store immediately at –80 °C to recombineer the target BAC with other targeting cassettes in the future.

Rhamnose induces the expression of the lambda phage proteins that carry out recombineering (Red αβγ) from the psc101-gbaA plasmid. The short period of growth at 37 °C assists with the expression of these proteins. After 40 minutes, the culture should be in mid-log phase and slightly cloudy.

Once the targeting cassette is introduced into the cells, a small number of the cells will replace the region between the homology arms with the targeting cassette. These cells are the rare recombinants and occur at a rate of 1 in 10,000–100,000 cells ( Datta et al., 2008 ).

Do not use a tetracycline resistance gene to select for recombinants, as psc101-gbaA already confers tetracycline resistance. If editing a BAC that already confers chloramphenicol resistance, do not use a chloramphenicol resistance gene to select for recombinants. In general, verify what antibiotics the target strains are already resistant to, because these antibiotics cannot be used to select for recombinants.

There should be a few dozen to a few hundred colonies on the plate. If there is a lawn or thousands of colonies, the targeting cassette was likely contaminated with template plasmid from the PCR. Any template plasmid that enters the target cells will confer antibiotic resistance without successful recombination. See Support Protocol 1 for details on how to purify the targeting cassette and remove template plasmid.

• 29. In order to screen for successful recombinants, use the sequencing primers from step 4 to perform colony PCR as described in Support Protocol 2 .

Targeting primers and targeting cassette PCR products often contain random mutations, so it is important to obtain full sequencing coverage of the final recombineering product to ensure it is free of undesired mutations ( Thomason et al., 2014 ). Order additional sequencing primers if necessary for large modifications, as most Sanger sequencing reads are less than 1000 bp. Ordering sequencing primers spaced out every 500 bp over a region of interest generally provides reliable coverage with overlapping reads. For example, a 5000 bp insertion should have about 8 equally spaced sequencing primers throughout the insertion in addition to the standard sequencing primers that flank the insertion.

The E. coli genome has its own dedicated system for resolving genome dimers involving FtsK and xerCD, which splits chromosome dimers back into monomers before the cell divides ( Bigot, Sivanathan, Possoz, Barre, & Cornet, 2007 ). Therefore, it is generally not necessary to check for heteromultimerization when modifying the E. coli genome. When modifying a BAC, colony PCR and sequencing can confirm that the modification is present, but they are generally unreliable techniques for identifying heteromultimers. Theoretically, heteromultimers should yield two different colony PCR bands on a gel with two different sizes, but in practice the smaller PCR product generally amplifies more efficiently than the larger product. Sometimes the larger product is not even visible, despite actually being present.

• 32. Use the successful recombinant to inoculate a 5 mL culture of LB with 5 μL of 1000X kanamycin stock solution in a sterile 14 mL culture tube. Incubate the culture overnight at 30 °C, shaking at 250 r.p.m. for ~16 hours.
• 33. Thoroughly mix 500 μL of the overnight culture with 500 μL of sterile 50% glycerol in a sterile 1.7 mL microcentrifuge tube (without a hole in the lid) and store immediately at –80 °C to recombineer the target BAC with other targeting cassettes in the future.
• 34. To isolate the target BAC DNA, follow steps 35–44 to perform isopropanol precipitation, as adapted from previous work ( Warming, Costantino, Court, Jenkins, & Copeland, 2005 ).
• 35. Pellet the rest of the overnight culture at 4,500 × g for 5 minutes. Then resuspend the pellet in 250 μL of Solution I from the Omega Bio-tek Plasmid Mini Kit and transfer to a 1.7 mL microcentrifuge tube.
• 36. Add 250 μL of Solution II from the Omega Bio-tek Plasmid Mini Kit to the resuspended cells, mix by inversion, and incubate at room temperature for 3 minutes exactly to lyse the cells. The mixture should appear more transparent after adding Solution II.
• 37. Add 250 μL of Solution III from the Omega Bio-tek Plasmid Mini Kit to the lysed cells and mix by inversion immediately to precipitate cellular debris, which appears as white chunks.
• 38. Incubate the lysed cells on ice for 5 minutes, then pellet the cellular debris at ≥16,000 × g for 5 minutes.
• 39. Transfer the clear supernatant to a new 1.7 mL microcentrifuge tube and pellet any remaining debris with a second spin at ≥16,000 × g for 5 minutes.
• 40. Transfer the clear supernatant to another new 1.7 mL microcentrifuge tube. Add 750 μL of isopropanol, mix thoroughly by inversion, and incubate on ice for 10 minutes. The solution should start to turn cloudy as the DNA precipitates.
• 41. Pellet the DNA at ≥16,000 × g for 10 minutes, then remove the supernatant without disturbing the white pellet at the bottom of the tube.

If the pellet becomes dislodged, spin at ≥16,000 × g for 1 minute to secure it back to the bottom of the tube.

• 43. Remove the 70% ethanol and air dry the pellet with the lid open for 10 minutes.
• 44. Resuspend the pellet in 50 μL of molecular biology-grade water.

Bear in mind that the psc101-gbaA plasmid has a copy number of about 10 plasmids per cell, and will also have a fragmentation pattern that will be much brighter and appear on top of the target BAC fragmentation pattern ( Fig. 4B ). Choose an enzyme that cuts the psc101-gbaA plasmid at least once, because uncut psc101-gbaA normally yields a complex pattern. If the chosen enzyme that cuts the target BAC does not cut psc101-gbaA, add a second enzyme to digest the psc101-gbaA plasmid. However, remember to account for how the second enzyme might affect the target BAC pattern.

Example strategy to identify heteromultimers using restriction enzyme digestion patterns. ( A ) Simplified plasmid maps are shown with the predicted sites of cleavage by the XbaI restriction endonuclease. XbaI is chosen in this example because it cleaves the psc101-gbaA recombineering plasmid at least once and because the desired modification (Mod) introduces a new XbaI site. ( B ) ( Left ) XbaI yields a predicted fragmentation pattern of the parent BAC that is distinct from that of the modified BAC. In particular, the parent BAC pattern has a unique band (red arrow) that is not present in the modified BAC pattern, and the modified BAC pattern has unique bands (blue arrows) that are not present in the parent BAC pattern. Furthermore, these unique, identifying bands are not obscured by the psc101-gbaA digestion pattern or other bands in the digestion patterns. The experimental gel image ( Right ) for this example shows that the hetermultimer contains both the unique parent BAC band (red arrow) and the unique modified BAC bands (blue arrows). The pure modified BAC pattern lacks the unique parent BAC band. The brightness and contrast in the gel image has been adjusted to render the faint bands more visible.

• 46. Digest 25 μL of the isolated target BAC DNA using the selected endonucleases following the manufacturer’s instructions. Additionally, digest 25 μL of the unmodified parent BAC as a negative control.

Preparing and running the agarose gel to obtain digestion patterns

Monitor the agarose while microwaving to ensure that it does not boil over. Use oven mitts or paper towels when handling the hot flask.

• 48. Add 5 μL of GelGreen dye to the molten agarose, swirl the flask to distribute the dye, then pour into a tray in a gel caster with an 8-well comb. Allow the gel to cool and solidify for 15–30 minutes.
• 49. Carefully remove the comb and place the solidified gel into an electrophoresis chamber filled with 0.5X TBE buffer. Ensure that there is sufficient TBE buffer to submerge the entire gel.
• 50. To check the digestion product, mix 10 μL of the digestion reaction with 2 μL of 6X gel loading dye, then load it into one of the wells of the agarose gel next to another well that contains 5 μL of Quick-Load Purple 1 kb DNA Ladder.

If the bands are not sufficiently separated, continue to run the gel for additional 10 minute increments until separation is sufficient.

• 52. If the fragmentation pattern unambiguously matches the expected pattern for the recombinant BAC and not the pattern for the unmodified BAC, return to step 23 to continue with further modifications as required. If there are no further modifications to make to the target BAC, see Support Protocol 3 to remove the psc101-gbaA plasmid from the cell. If the fragmentation pattern corresponds to the expected pattern for the recombinant BAC overlaid on top of the pattern for the unmodified BAC, or if there is any doubt, proceed to Support Protocol 4 to “remonomerize” the target BAC and obtain a pure monomer of the desired recombinant BAC.

Because of the low copy number of BACs per cell, it can sometimes be quite difficult to isolate enough DNA to analyze by gel electrophoresis. If the bands of the restriction digestion pattern are too faint to confidently identify successful recombinants or heteromultimers, grow a 25 mL culture of each target BAC in question and isolate the DNA using the ZymoPURE II Plasmid Midiprep Kit D4200, following the manufacturer’s instructions. Midiprep kits, while more expensive, isolate a larger quantity of DNA, which can make visualizing digestion patterns easier when isopropanol precipitation is insufficient.

BASIC PROTOCOL 2

Direx recombineering for seamless deletions or small insertions.

If many sequential modifications need to be made in the target or, as is the case for point mutations, the modification must be made without leaving an antibiotic resistance gene behind, then Basic Protocol 2 or 3 should be used. Further, if the modification is a point mutation, deletion, or a small insertion/replacement (≤30 bp), DIRex recombineering is the most rapid approach to introduce these modifications ( Näsvall, 2017 ). First, a kanamycin resistance gene ( kan R ) and a ccdB gene, which is a counterselection gene that is lethal only under certain conditions, replace or are inserted into the target region that will be modified ( Fig. 5A ). Subsequently, k an R + ccdB -containing intermediates are selected for using kanamycin and arabinose, which induces expression of the ccdA antitoxin that renders ccdB non-lethal ( H. Wang et al., 2014 ). The kan R and ccdB genes are flanked by inverted repeats, which are in turn flanked by homology arms that contain a direct 30 bp repeat centered around the modification. The architecture of the indirect and direct repeats promotes spontaneous excision of the kan R gene, the ccdB gene, the inverted repeats, and one of the direct repeats, thereby leaving behind a single copy of the modification ( Fig. 5B ). The excision event is thought to occur through strand slippage and mispairing during replication ( Bzymek & Lovett, 2001 ; Näsvall, 2017 ). Successful recombinants are then selected for by removing arabinose such that ccdA is no longer expressed and the ccdB toxin becomes lethal. Any cells that have not excised ccdB to yield the final recombinant, which will be the vast majority of cells, will not survive this selection pressure. We note that there are many potential options for counterselection genes available, but the protocols in this article utilize ccdB because in our experience it functions quickly and reliably and without requiring the use of minimal media plates that slow the growth of cells. Finally, we note that although various scales can be used, we strongly recommend (and describe below) performing the protocol on a 1 mL scale for ease of sample processing and using heat blocks, as described in previous work ( H. Wang et al., 2014 ).

Schematic of DIRex recombineering. ( A ) In DIRex recombineering, intermediates containing the conditionally-lethal gene ( ccdB ) are first selected for using kanamycin. The modification (Mod) is directly repeated in the homology arms. Thus, the direct and inverted repeats (IR) promote spontaneous excision to yield the final recombinant. Successful recombinants are selected for by removing arabinose. In the absence of arabinose, the ccdA antitoxin is no longer expressed and ccdB then kills unmodified cells. ( B ) DIRex is hypothesized to promote spontaneous excision through hybridization between the two inverted repeats to form a hairpin during replication that brings the direct repeats into close proximity ( Bzymek & Lovett, 2001 ; Näsvall, 2017 ). The direct repeats can then promote strand slippage during synthesis that results in excision of one of the direct repeats and everything between the direct repeats.

R6K-KCA plasmid (see Table 1 )

R6K-AKC plasmid (see Table 1 )

Lysogeny broth media (see recipe)

Super optimal broth with catabolic repression media (see recipe)

10% arabinose stock solution (see recipe)

LB agar plates with 1X kanamycin and 0.2% w/v arabinose (see recipe)

30 °C incubator (e.g., VWR Gravity convection incubator 89511–422)

Primer design example for a small seamless insertion via DIRex recombineering. ( A ) The left and right targeting primers are used to amplify the left and right targeting half-cassettes from the R6K-AKC and R6K-KCA template plasmids ( Table 1 ), respectively. The overhangs of the left and right targeting cassettes contain the left (red) or right (blue) homology arms that flank the target region to be replaced, respectively, along with a direct repeat that contains the small insertion (Mod) and a small portion of the other homology arm. ( B ) The left homology arm is copied from the top strand to the left of the target region (red color), and the right homology is copied from the bottom strand to the right of the target region (blue color). ( C ) A simple strategy to design the full homology arms with the direct repeat is to generate the desired sequence with the modification (brown) between the left and right homology arms from B . The full homology arm of the left targeting primer is shown as the underlined region of the top strand and the full homology arm of the right targeting primer is shown as the underlined region of the bottom strand. The 3′ ends of these full homology arms overlap by 30 bp, with the modification in the center of the overlap. In other words, each homology arm contains the modification on the 3′ end followed by N bases of the complement of the other homology arm, where N = 15 – ((the size of the insertion)/2). ( D ) Finally, the full left and right homology arms are appended to the 5′ ends of the DIRexPrimer sequence ( Table 2 ) to form the full left and right targeting primers.

Guide for homology arm design. ( A ) Homology arms (red and blue) are introduced as 5′ overhangs on primers (green boxes) that amplify the targeting cassette via PCR. ( B ) Homology arm overhangs for insertions, point mutations, deletions, and replacements are highlighted as green boxes. The modification targeting cassettes for point mutations and deletions using two-step recombineering ( Basic Protocol 3 ) are small and can be generated by annealing oligos rather than by PCR.

• A deletion: Take the reverse complement of the last 15 bp of the right homology arm and append it to the 3′ end of the left homology arm. Take the reverse complement of the last 15 bp of the left homology arm—not including the bases appended from the right homology arm—and append it to the 3′ end of the right homology arm. The resulting 65 bp homology arms complement each other for the last 30 bp of their 3′ ends, which will generate a 30 bp direct repeat of the deletion junction near each end of the targeting cassette. Append each of these homology arms to the DIRexPrimer sequence in Table 2 to create the left and right targeting primers.

Note that the target region to be replaced can be large or small, it is only the new insertion that is replacing the target region that is limited to ~30 bp. The size of the insertion is only limited by the size of the DNA oligos that can be purchased. Most vendors currently limit standard oligo sizes to 100 or 120 bp.

Order the left and right targeting primers, then use them to create the targeting half-cassettes. Generate the left half-cassette using the left targeting primer and DIRexKan.Rev primer from Table 2 in conjunction with R6K-AKC as the PCR template (see Support Protocol 1 ). Generate the right half-cassette using the right targeting primer and the DIRexKan.For primer from Table 2 in conjunction with R6K-KCA as the PCR template (see Support Protocol 1 ).

Because the direct repeats in the overhangs of the left and right targeting primers need to flank an inverted repeat, the inverted repeat-binding site at the 3′ end of the left and right target primers is identical. If the left and right targeting primers were used simultaneously in the same PCR reaction to generate the targeting cassette, there would be a mixture of PCR products that have only the left homology arms, only the right homology arms, or both homology arms. By designing the targeting cassette with slightly overlapping halves, each targeting cassette will necessarily contain one left homology arm and one right homology arm with a defined orientation. The half-cassettes will be electroporated together and joined by homologous recombination inside the cell ( Fig. 5A ).

• Design and order sequencing primers as in Basic Protocol 1 , step 4 that are 200 bp to the left of the left homology arm and 200 bp to the right of the right homology arm.
• 5. If the target BAC and the psc101-gbaA plasmid are already both present in DH10B cells, or if the intent is to engineer the E. coli genome and the psc101-gbaA plasmid is already present in the strain to be engineered, proceed to step 6. Otherwise, carry out Basic Protocol 1 , steps 5–22.

Note, when engineering the E. coli genome, omit the addition of chloramphenicol to all agar plates and media in this step and all subsequent steps.

Rhamnose induces the expression of the lambda phage proteins that carry out recombineering (Red αβγ) from the psc101-gbaA plasmid. The arabinose induces the ccdA antitoxin that binds the ccdB toxin and renders it non-lethal. The short period of growth at 37 °C assists the expression of these proteins. After 40 minutes, the culture should be in mid-log phase and slightly cloudy.

Do not forget to add arabinose to ensure that ccdB does not kill the cells.

• 10. After the 2-hour recovery period, pellet the cells at 10,000 × g for 1 minute. Resuspend the pellet in 50 μL of LB media.

There should be a few dozen to ~100 colonies on the plate.

Because there is no arabinose on the plate and thus no expression of the ccdA antitoxin, only those rare cells that have undergone spontaneous excision of ccdB will survive. Split the plate into thirds and streak each of the 3 colonies in a zig-zag pattern across a third of the plate. Three colonies are picked because 30–50% of colonies will be heteromultimers of unmodified parent and kan R +ccdB intermediate BACs. Heteromultimers easily undergo intramolecular recombination to yield parent BAC and survive counterselection ( Fig. 8 ) at a frequency much higher than spontaneous excision. Therefore, it is important to pick several colonies to increase the chances of choosing a monomer. Note that it is simpler and faster to just streak several colonies rather than to screen for monomers of the kan R +ccdB intermediate. Open in a separate window Figure 8 Schematic showing the outcomes of recombineering with monomeric kan R + ccdB intermediates (left) compared to recombineering with heteromultimers of kan R + ccdB intermediate and unmodified parent (right). The monomeric intermediate must undergo recombineering with the targeting cassette to eliminate the toxic ccdB gene. However, the heteromultimer can undergo intramolecular recombination to eliminate the toxic ccdB gene, yielding unmodified parent BAC at a frequency much higher than successful recombineering. Ensure that chloramphenicol is present in the plates. Without chloramphenicol, cells can survive counterselection by losing the ccdB-containing target BAC, which happens at a frequency much greater than the frequency of successful recombination. Chloramphenicol will kill any cells that lose the target BAC.

When a monomer kan R +ccdB intermediate is streaked on a plate in the absence of arabinose, a dozen or a few dozen isolated colonies should appear. These colonies will have undergone spontaneous excision to yield the final recombinant. If a heteromultimer is streaked on a plate, a lawn of bacteria will grow. These bacteria have recombined to yield unmodified parent BAC. In further steps, only screen colonies from streaks that have yielded just a few dozen isolated colonies.

If every colony chosen, rather than just 30–50% of the colonies, behaves as a heteromultimer, it could be that the parent target BAC was a homomultimer prior to the insertion of the kan R +ccdB cassette ( Fig. 9 ). In such a situation, nearly 100% of kan R +ccdB intermediates will be heteromultimers. To address this issue, the target BAC must be remonomerized using Support Protocol 4 before reattempting this protocol. Open in a separate window Figure 9 Schematic showing the outcome of recombineering with a homomultimer. Target BACs that are homomultimeric will almost always form heteromultimeric kan R + ccdB intermediates, which are unsuitable for counterselection owing to the high rate of intramolecular recombination to yield unmodified parent BAC.

• 15. In order to screen for successful recombinants, use the sequencing primers from step 4 to perform colony PCR as described in Support Protocol 2 .

Targeting primers and targeting cassette PCR products often contain random mutations, so it is important to get full sequencing coverage of the final recombineering product to ensure it is free of undesired mutations ( Thomason et al., 2014 ). Order additional sequencing primers if necessary for large modifications as most Sanger sequencing reads are less than 1000 bp. Ordering sequencing primers spaced out every 500 bp over a region of interest generally provides reliable coverage with overlapping reads. For example, a 5000 bp insertion should have about 8 equally spaced sequencing primers throughout the insertion in addition to the standard sequencing primers that flank the insertion.

• 17. After sequence confirmation, return to step 5 to introduce further modifications as needed. If there are no additional modifications to make, see Support Protocol 3 to remove the psc101-gbaA plasmid from the cell.

We note that during counterselection the toxic ccdB gene selects against any potential heteromultimers that form via fusion of a kan R +ccdB intermediate BAC and successfully modified recombinant BAC. Heteromultimers of unmodified parent BAC and a kan R +ccdB intermediate BAC almost always yield parent BAC after counterselection, so a heteromultimer of unmodified parent BAC and successfully modified recombinant BAC is very unlikely. Therefore, unlike in Basic Protocol 1 , it is unlikely that the final recombinant is a heteromultimer. Hence, it is generally not necessary to screen for heteromultimers. If in doubt, carry out Basic Protocol 1 , steps 32–52 to ensure that the final recombinant BAC is not a heteromultimer.

BASIC PROTOCOL 3

Two-step recombineering for large, seamless insertions or for gene replacements.

For large (>30 bp), seamless insertions or replacements, a standard two-step recombineering protocol should be used ( H. Wang et al., 2014 ). We note that two-step recombineering can also be used to make deletions, point mutations and small insertions/replacements, but it is an unnecessarily laborious way to do so compared to Basic Protocol 2 . For the first step of Basic Protocol 3 , a kan R gene and a ccdB gene replace or are inserted into the target region that will be modified ( Fig. 10 ). Kan R + ccdB -containing intermediates are selected for by streaking on plates containing kanamycin and arabinose, which induces expression of the ccdA antitoxin that renders ccdB non-lethal. In the second step, the kan R and ccdB genes in the intermediate are replaced by a targeting cassette that introduces the final modification. Successful recombinants are selected for by removing arabinose, such that ccdB is rendered lethal and unmodified cells do not survive ( H. Wang et al., 2014 ). We note that there are many potential options for counterselection genes available. The protocols in this article utilize ccdB because, in our experience, it functions quickly and reliably and without requiring the use of minimal media plates that slow the growth of cells. Finally, we note that although various scales can be used, we strongly recommend (and describe below) performing the protocol on a 1 mL scale for ease of sample processing and using heat blocks, as described in previous work ( H. Wang et al., 2014 ).

Schematic of two-step recombineering. First, an intermediate containing a conditionally lethal gene ( ccdB ) is selected for using kanamycin. Second, the ccdB gene in the intermediate is replaced with the desired modification, which is selected for by removing arabinose. In the absence of arabinose, the ccdA antitoxin is no longer expressed and ccdB kills unmodified intermediate cells, leaving only the successful recombinant cells that have the final modification.

Template plasmid (for insertions, supplied by user)

• First, choose appropriate homology arms to make the desired modification. Immediately to the left of the region to be deleted or replaced, highlight 50 bp, not including the region to be deleted or replaced, and copy the sequence from the top strand (in the 5′ to 3′ direction) for the left homology arm.
• Immediately to the right of the region to be deleted or replaced, highlight 50 bp, not including the region to be deleted or replaced, and copy or mark the sequence from the BOTTOM strand for the right homology arm. Ensure that the sequence copied and designated as the right homology arm is the bottom strand in the 5′ to 3′ direction (i.e., the reverse complement of the TOP strand). In the case of an insertion, the 3′ ends of the left homology arm and the right homology arm will be adjacent to each other if the homology arms are properly designed. In the case of a point mutation, the base to be modified should lie between the 3′ ends of the homology arms ( Fig. 7 ).

In the case of deletions, the modification targeting cassettes for the second step are so small that they are most easily made by ordering the entire top and bottom strands of the cassette as oligos and annealing them according to Support Protocol 1 .

• A large >30 bp insertion or replacement: Create or obtain a plasmid that has the genes or region to be inserted into the target. Design primers that have melting temperatures ( T m ) of ~60 °C and that amplify the entire region to be inserted ( Sawitzke et al., 2013 ). In the primer design, append the left homology arm to one primer and the right homology arm to the other primer. Note that the choice of which homology arm to append to which primer will determine the ultimate orientation of the insertion. Once the primers are obtained, use the primers in conjunction with the template plasmid to create the targeting cassette via PCR (see Support Protocol 1 ).
• A point mutation or a small <30 bp insertion: Append the modified base or small insertion to the 3′ end of the left homology arm sequence, followed by the reverse complement of the right homology arm sequence to create the forward modification oligo. Append the reverse complement of the modified base or small insertion to the right homology arm sequence, followed by the reverse complement of the left homology arm sequence to create the reverse modification oligo. Anneal the forward and reverse modification oligos to generate the modification targeting cassette according to Support Protocol 1 .

In the case of point mutations or small insertions, the modification targeting cassettes for the second step are so small that they are most easily made by ordering the entire top and bottom strands of the cassette as oligos and annealing them. The size of the homology arms can be reduced to meet oligo manufacturing size limits, but can be no smaller than 35 bp. Bear in mind that smaller homology arms will result in reduced recombineering efficiency ( Yu et al., 2000 ). Also, be sure to double-check the orientation of the insertion.

Recombineering with the targeting cassettes

• 5. If the target BAC and the psc101-gbaA plasmid are already both present in DH10B cells, or if the intent is to engineer the E. coli genome and the psc101-gbaA plasmid is already present in the strain to be engineered, proceed to step 6. Otherwise, perform Basic Protocol 1 , steps 5–22.

Note that, if engineering the E. coli genome, omit the addition of chloramphenicol to all agar plates and media in this step and all subsequent steps.

Rhamnose induces the expression of the lambda phage proteins that carry out recombineering (Red αβγ) from the psc101-gbaA plasmid. The arabinose induces the ccdA antitoxin that binds the ccdB toxin and renders it non-lethal. The short period of growth at 37 °C assists the expression of these proteins. After the 40 minutes, the culture should be in mid-log phase and slightly cloudy.

Arabinose needs to always be present at concentration of 0.2% v/v in the media or in agar plates from this point forward to ensure that ccdB does not kill the cells. If you forget to add arabinose in any of the subsequent steps, with the exception of the molecular biology-grade water washes before the second electroporation, you must restart the protocol. While the culture may eventually grow in the absence of arabinose, it will consist of cells that have inactivated ccdB via random mutations, thus rendering counterselection impossible in the second step. Do not omit arabinose until explicitly instructed to do so at the end of the protocol.

Three colonies are picked because 30–50% of colonies will be heteromultimers of unmodified parent and kan R +ccdB intermediate BACs. Heteromultimers easily undergo intramolecular recombination to yield parent BAC to survive counterselection ( Fig. 8 ) at a frequency much higher than successful recombineering. Therefore, it is important to pick several colonies to increase the chances of choosing a monomer. Note that it is simpler and faster to just pick several colonies to recombineer in the second step rather than to screen for monomers of the kan R +ccdB intermediate.

Optionally, thoroughly mix 500 μL of each overnight culture with 500 μL of sterile 50% glycerol in a sterile 1.7 mL microcentrifuge tube (without a hole in the lid) and store immediately at –80°C. Once a kan R +ccdB intermediate is confirmed to be monomeric via successful recombineering, it can be used again in the future to introduce modifications into the same location. This intermediate can be used to make any other modification in the region as long as the homology arms straddle the kan R +ccdB genes. Furthermore, when generating a set of different modifications in the same location, we note that it is useful to pilot just one modification of a set to obtain a confirmed kan R +ccdB intermediate monomer. The confirmed monomer can then be used to complete the set of modifications in parallel rather than using three different intermediates. This approach reduces the number of samples that need to be processed by a factor of three.

Rhamnose induces the lambda phage proteins that carry out recombineering. Arabinose should already be present in the culture to maintain ccdA antitoxin expression.

It is particularly important that the recovery time is at least 2 hours at this step so that the cells divide enough times to isolate successful recombinants from any unmodified kan R +ccdB intermediates ( Warming et al., 2005 ). During mid-log phase, there are usually the equivalent of ~4–8 copies of the genome, and presumably of BACs as well, in a single cell ( Åkerlund, Nordström, & Bernander, 1995 ). It is likely that only one of these will be modified in a cell where successful recombination has occurred. Only one copy of the ccdB gene is required to kill a cell. Thus, it is critical that any successful recombinant genomes or BACs have time to segregate into daughter cells away from any copies of ccdB before arabinose is removed and ccdA antitoxin production ceases.

It is important to remove the arabinose from the cells in this step by pelleting and resuspending in 1 mL LB media. In the absence of arabinose, ccdB will kill any cells that still carry the gene, thus counterselecting against unmodified cells and selecting for successful recombinants.

Do not plate all the cells from the 1 mL culture onto the plate. The ccdB gene does not kill cells immediately and the cells will divide for several rounds before dying ( Jaffé, Ogura, & Hiraga, 1985 ). If too many cells are plated at once, they will grow into a lawn before dying. This lawn obscures successful recombinants and prevents them from forming distinguishable colonies. 50 μL out of the 1 mL culture (1/20 th of the cells) on one half of a 10 cm plate generally yields good results, but sometimes still forms a lawn. Therefore, it is also useful to plate a more dilute sample on the other half of the plate. Ensure that chloramphenicol is present in the plates. Without chloramphenicol, cells can survive counterselection by losing the ccdB-containing target BAC, which occurs at a frequency much greater than the frequency of successful recombination. Chloramphenicol will kill any cells that lose the target BAC.

Either the more concentrated or more dilute half of the plate should have a few dozen distinguishable colonies. Plates with hundreds or thousands of colonies are likely the result of heteromultimers that recombined to yield unmodified parent BAC and are thus not worth screening. If, after attempting the protocol a few times, every plate consistently has hundreds or thousands of colonies, it could be that the parent target BAC was a homomultimer prior to the insertion of the kan R +ccdB cassette ( Fig. 9 ). In such a situation, nearly 100% of kan R +ccdB intermediates will be heteromultimers, rather than just 30–50%. To address this issue, the target BAC must be remonomerized using Support Protocol 4 before reattempting this protocol.

Insertions of ~2 kb and larger undergo recombineering quite inefficiently ( Kuhlman & Cox, 2010 ). In such a case, many of the colonies on the plate will be false positives in which the ccdB gene has acquired a mutation that renders it inactive and/or non-lethal. Inactivation of ccdB is an event that occurs at a frequency similar to recombineering events, so it is especially important to screen many colonies (24–96 colonies) when recombineering with large cassettes.

• 22. After sequence confirmation, return to step 5 to introduce further modifications as needed. If there are no additional modifications to make, see Support Protocol 3 to remove the psc101-gbaA plasmid from the cell.

The glycerol stock of the kan R +ccdB intermediate from step 14 that yielded a successful recombinant should be saved as a “confirmed monomer”. We note that during counterselection the toxic ccdB gene selects against any potential heteromultimers that form via fusion of a kan R +ccdB intermediate BAC and successfully modified recombinant BAC. Heteromultimers of unmodified parent BAC and a kan R +ccdB intermediate BAC almost always yield parent BAC after counterselection, so a heteromultimer of unmodified parent BAC and successfully modified recombinant BAC is very unlikely. Therefore, unlike in Basic Protocol 1 , it is unlikely that the final recombinant is a heteromultimer. Hence, it is generally not necessary to screen for heteromultimers. If in doubt, carry out Basic Protocol 1 , steps 32–52 to ensure that the final recombinant BAC is not a heteromultimer.

ALTERNATE PROTOCOL 1

Larger scale cultures for inefficient recombineering.

Basic Protocols 1 – 3 use 1 mL of mid-log cultures to generate electrocompetent cells. This scale is convenient to process and is generally sufficient for routine recombineering ( H. Wang et al., 2014 ). However, increasing the number of electrocompetent cells for difficult, inefficient recombineerings—such as for large inserts or replacements—can increase the chances of obtaining a successful recombinant when 1 mL scale recombineering has failed. This protocol describes how to perform Basic Protocol 3 on a 25 mL scale rather than a 1 mL scale, as adapted from previous work ( Warming et al., 2005 ).

Target BAC-kanR+ccdB intermediate (from Basic Protocol 3 , steps 6–13)

Modification targeting cassette (from Basic Protocol 3 , step 3)

Sterile 10 μL pipette tips (VWR 89079–466)

Sterile 200 μL pipette tips (VWR 89079–460)

Sterile 1000 μL pipette tips (VWR 89079–472)

10 mL serological pipette (Cellstar 607180)

Sterile 50 mL baffled flasks (Wheaton 354235)

Aluminum foil

50 mL conical tubes (Cellstar 227261)

Molecular biology-grade water (Corning 46-000-CM)

0.1 cm electroporation cuvettes (Bio-Rad 1652083)

14 mL sterile culture tubes (VWR 60818-703)

15 mL conical tube (Cellstar 188271)

DNA Editing Software (e.g Snapgene, ApE, Serial Cloner)

P10 pipettor (e.g. VWR 89079-962)

P200 pipettor (e.g. VWR 89079-970)

P1000 pipettor (e.g. VWR 89079-974)

Pipette gun (e.g. Drummond Portable Pipet-Aid XP 4-000-101)

30 °C shaking incubator (e.g. Thermo Fisher MaxQ 4000 SHKE4000)

UV-Vis spectrometer (e.g. Thermo Fisher Nanodrop 2000c)

Shaking water bath (e.g. Shel Lab SWBR17)

Refrigerated centrifuge capable of 4,500 × g with rotor and adapters for 15 mL and 50 mL conical tubes (e.g. Beckman Coulter Avanti J-E 369003 with JA-10 rotor 369687)

Refrigerated microcentrifuge (e.g. Thermo Fisher Heraeus Fresco 21 75002426)

Recombineering on a 25 mL scale

• Design and create targeting primers and cassettes as described in Basic Protocol 3 , steps 1–4.
• If target BAC and the psc101-gbaA plasmid are already both present in DH10B cells, or if the intent is to engineer the E. coli genome and the psc101-gbaA plasmid is already in the strain to be engineered, proceed to step 6. Otherwise, carry out Basic Protocol 1 , steps 5–22.

The insertion of the kan R +ccdB cassette is usually efficient enough to carry out on a 1 mL scale rather than a 25 mL scale. Ideally, perform large scale recombineering with a confirmed kan R +ccdB intermediate monomer when possible to reduce the number of large-volume samples that need to be processed. Otherwise, perform this protocol with three kan R +ccdB intermediates if there is no confirmed monomer available.

Sterilize the 50 mL glass baffled flasks by rinsing thoroughly with deionized water, covering the opening with aluminum foil to serve as a cap, and autoclaving.

Air incubators do not efficiently heat larger cultures in short periods of time, so it is important to use a shaking water bath. Alternatively, if a shaking water bath is not available the flasks can be swirled by hand in a 37 °C bath.

• After incubation at 37 °C, the culture should have an OD 600 of ~0.6. Place the culture on ice to arrest growth. Next, transfer the cells to a 50 mL conical tube and pellet at 4,500 × g in a refrigerated centrifuge at 4 °C for 5 minutes. Carefully remove the media with a 20 mL pipette to avoid disturbing the pellet.
• Gently resuspend the pellet in 10 mL ice-cold molecular biology-grade water by gently pipetting several times. Next, pellet again at 4,500 × g in a refrigerated centrifuge at 4 °C for 5 minutes.

After the first wash, the pellet becomes very loose and easy to disturb. Slowly remove media with the pipette gun from the top down. Do not pour off the media because this risks losing a significant quantity of the cells.

Often there is ~1 mL of residual water around the pellet in the bottom of the Falcon tube, in which case it is unnecessary to add additional water. In the next step, the cells are concentrated further.

• Pellet the cells at 10,000 × g in a refrigerated microcentrifuge at 4 °C for 1 minute.

Often there is ~20 μL of residual water around the pellet, in which case it is unnecessary to add additional water to the tube. The resuspended cells should appear opaque. Because this batch of electrocompetent cells will be more concentrated, ensure there is enough water to thoroughly resuspend the pellet with no clumps.

• Add ~200 ng of the modification targeting cassette to the electrocompetent cells and mix gently by pipetting a few times. Keep the tube on ice.
• Before electroporating, set up all of the necessary electroporation materials. Plug in the Bio-Rad micropulser and turn it on. The screen should display the default setting “Ec1”, which is the proper setting to use. Press the measurement button twice until a light appears next to the “ms” label and the screen reads “0.00”. Set a bottle of SOC media close to the electroporator with the cap completely loose for quick access. Ensure that a box of Kimwipes is nearby.
• Add the electrocompetent cells that have been mixed with DNA into the 0.1 cm gap between the metal plates of a pre-chilled electrocuvette. Next, quickly place the cap back onto the electrocuvette.
• Wipe any condensation or ice off the outside of the electrocuvette, then place it into the sample holder with the notch facing forward. Gently slide the electrocuvette into the electroporation chamber until the electrocuvette “clicks” between the flexible metal electrodes in the electroporation chamber.

If the pulse length is less than 4.80 ms or if the screen says “Arc”, repeat the electroporation with new electrocompetent cells and add a smaller quantity of DNA. It may be necessary to desalt the DNA again or wash the cells an extra time. It is often useful to prepare a few batches of electrocompetent cells in parallel in case the electroporation needs to be repeated a few times before it is successful.

• Add the resuspended cells to 4 mL of SOC media in a sterile 14 mL culture tube and add 100 μL of 10% arabinose stock solution. Incubate the culture at 30 °C for 2 hours at 250 r.p.m in a shaking incubator.
• After recovery, pellet the cells in a 15 mL conical tube at 4,500 × g for 5 minute. Resuspend the pellet in 1 mL of LB media.
• Carry out Basic Protocol 3 , steps 18–22.

SUPPORT PROTOCOL 1

Creating targeting cassettes.

The quality of targeting cassette DNA is paramount to the success of recombineering. It is essential to obtain low-salt, high-concentration DNA samples that are relatively free of PCR side products or other unwanted DNA, such as template plasmids that can result in false positives upon selection. Cassettes of 100 bp or smaller can easily be generated by purchasing the top and bottom strand as separate oligos and annealing them to yield cassettes with high-concentration and purity, as described in previous work ( Warming et al., 2005 ). Targeting cassettes too large to order as oligos can be generated by PCR using the targeting primers and a PCR template. However, it is important to be aware if two important issues when generating targeting cassettes by PCR:

• PCR often generates a series of truncated side products from non-specific primer binding. These side products are sometimes, but not always, visible on an agarose gel as faint bands or smears below the expected band. Because smaller targeting cassettes recombine with a much higher efficiency than larger cassettes, these truncated side products often recombine into the region of interest much more often than the main desired cassette. This problem is particularly significant when the desired cassette is very large. Therefore, it is wise to carefully purify cassettes larger than 2 kb via gel extraction to remove these truncation products.
• In cases where the PCR template plasmid encodes an antibiotic resistance gene or some other gene that will be applied to select for successful recombinants, it is absolutely critical to inactivate or completely remove the PCR template plasmid from the final targeting cassette product. If any intact PCR template plasmid enters the cells that are being recombineered, it will result in colonies after selection that have not actually undergone recombination. These false positives have simply taken up template plasmid leftover from the PCR reaction. If there is even minimal intact template plasmid contamination, these ‘cheaters’ will greatly outnumber any rare, successful recombinants. The most efficient approach to prevent false positives resulting from template plasmid uptake is to use DNA templates that cannot replicate in DH10B E. coli . Otherwise, the PCR product must be fully digested with endonucleases that destroy only the template plasmid, and/or be purified by gel extraction.

Targeting primers or oligos (designed and ordered by user)

Molecular biology-grade water (e.g., Corning 46–000-CM)

Q5 Reaction Buffer (New England Biolabs B9027S)

Lid lock (e.g., VWR 14229–941)

3 M sodium acetate (see recipe)

100% ethanol (e.g., Koptec V1001)

8-well PCR tube strips (e.g., VWR 20170–004)

OneTaq Quick-Load 2X Master Mix with Standard Buffer (New England Biolabs M0486L)

10X Cutsmart Buffer (New England Biolabs B7204S)

DpnI endonuclease (New England Biolabs R0176S)

Industrial razor blades (e.g., VWR 55411–050)

Gel extraction kit (e.g., Omega Bio-tek Gel Extraction Kit D2500–01)

PCR clean-up kit (e.g., Omega Bio-tek Cycle Pure Kit D6492–01)

Refrigerated microcentrifuge (e.g., Thermo Fisher Heraeus Fresco 21 75002426)

500 mL glass beaker (e.g., Corning 1003–600)

Magnetic stir bar (e.g., VWR 58947–132)

Foam float for 1.7 mL tubes (e.g., VWR 82017–634)

Stirring hot plate with temperature probe (e.g., Corning PC-420D 6795–420KIT)

Thermal cycler (e.g. Bio-Rad T100 Thermal Cycler 1861096)

Precision balance (e.g. Mettler Toledo ME1002TE 30216559)

Agarose gel electrophoresis system (e.g. Bio-Rad Mini-Sub Cell GT System 1664401)

37 °C incubator (e.g., VWR Gravity convection incubator 89511–422)

60 °C heat block (e.g., Corning LSE Digital Dry Bath 6875-SB with 24 × 1.5 mL block 480119)

Annealing oligos to generate small targeting cassettes ≤100bp

• Order oligos as desalted 100 μM stock solutions or as dry desalted oligos. Dissolve dry desalted oligos in molecular biology-grade water to yield 100 μM stock solutions. Be sure to thoroughly mix by vortexing the capped tube for ~1 minute, followed by spinning the tube at 10,000 × g for 1 minute.
• Add the following reagents to a sterile 1.7 mL microcentrifuge tube: 70 μL of molecular biology-grade water, 20 μL of 5X Q5 Reaction Buffer, 3 μL of 100 μM top strand oligo stock solution, and 3 μL of 100 μM bottom strand oligo stock solution. Close the tube and secure the lid with a lid lock.

Cover the beaker with an aluminum foil lid to prevent heat loss and accelerate heating.

• Place the tube from step 2 in a foam float and float it on the boiling water for 5 minutes.

For proper annealing, it is important to cool the samples slowly from boiling to room temperature over the course of at least 30 minutes. Do not just boil the tube then place it at room temperature. Use a sufficient volume of water to ensure slow cooling.

The solution should quickly become slightly cloudy as a precipitate forms.

A white pellet should be clearly visible at the bottom of the tube after centrifugation. This pellet contains the annealed oligos.

• Remove the supernatant, being careful to not disturb the white pellet at the bottom of the tube.

The pellet may dislodge, so be extra careful to not aspirate and discard the pellet.

This extra spin shifts residual ethanol and any dislodged pieces of the pellet back down to the bottom of the tube.

• Carefully remove any residual ethanol from around the pellet with a 10 μL pipette. Allow the contents of the open tube to air dry for ~10 minutes.
• Resuspend the pellet in 100 μL of molecular biology-grade water to obtain a targeting cassette solution. The resulting DNA concentration will be about ~200 ng/μL.

Generating large targeting cassettes by PCR

Dilute a portion of the 100 μM primer stocks to 10 μM before adding to the PCR reaction. Also, only add a very small amount of template DNA (~1 ng) so that it is easier to remove the template in later steps.

• Calculate the annealing temperature for the primers using the New England Biolabs Tm Calculator ( https://tmcalculator.neb.com/#!/main ). Ensure the polymerase option is set to “Q5 Hi-Fidelity 2X Master Mix”. Determine the expected size of the PCR product using DNA editing software.
• 98 °C        30 seconds
• 98 °C        10 seconds
• Annealing Temp.    30 seconds
• 72 °C        30 seconds per kb of expected PCR product
• Return to step 2 for 35 cycles
• 72 °C        2 minutes
• 4 °C          hold

GelGreen can be visualized using a blue LED transilluminator, which does not damage DNA. If possible, do not use ethidium bromide dye to visualize the PCR products, because it must be visualized using an ultraviolet transilluminator. Ultraviolet light will cause damage to the DNA products and increase the incidence of unwanted mutations.

• Carefully remove the comb and place the solidified gel into an electrophoresis chamber filled with 0.5X TBE buffer. Ensure that there is sufficient buffer to submerge the entire gel.
• To check the PCR product, mix 3 μL of PCR reaction with 1 μL of 6X gel loading dye. Next, load the resulting solution into one of the wells of the agarose gel next to another well that has 5 μL of Quick-Load Purple 1 kb DNA Ladder.
• Run the gel at 150 volts for 20 minutes, then visualize on a blue LED transilluminator.

If none of the temperatures in the annealing temperature gradient work, redo the gradient, but replace 10 μL of molecular biology-grade water with 10 μL of Q5 High GC Enhancer per reaction. Alternatively, try extending the 72 °C extension step to 60 seconds per kb of expected PCR product. If the PCR reaction still fails, try redesigning the primers to minimize primer-dimer formation. Visit https://www.thermofisher.com/us/en/home/brands/thermo-scientific/molecular-biology/molecular-biology-learning-center/molecular-biology-resource-library/thermo-scientific-web-tools/multiple-primer-analyzer.html to predict primer-dimer propensity.

As a general rule, use a PCR clean-up kit, such as the Omega Bio-tek Cycle Pure Kit, to purify the DNA whenever possible, instead of purifying DNA from the PCR reaction by gel extraction. Gel extraction generally provides very poor DNA yields, so it should only be used when absolutely necessary. Basic Protocols 1 – 3 use R6K-kan-ccdB, R6K-AKC, or R6K-KCA as template plasmids. These plasmids only have R6K origins that cannot replicate in DH10B cells and can only replicate in cells expressing the pi protein ( Kolter, Inuzuka, & Helinski, 1978 ), such as Pir2 E. coli cells. If the <2 kb PCR cassette was amplified from one of these three R6K plasmids, purify using the Omega Bio-tek Cycle Pure Kit. When there is a choice, always choose or design template vectors that are replication-incompetent in the strain to be used for recombineering. The R6K origin provides one simple and commonly applied option ( J. Wang et al., 2006 ).

E. coli that have an intact dam methylase methylate the adenine of every GATC sequence that occurs inside the cell. GATC is a very common sequence that occurs multiple times in most plasmids. DpnI is an endonuclease that can cut GATC only when the adenine is methylated. Adding DpnI digests the E. coli-derived template plasmid but leaves the newly synthesized, unmethylated PCR product intact. DpnI digestion will not be successful if the template is not methylated, such as when it is derived from E. coli that have lost dam methylase function, as indicated by the letters “dam” in the strain genotype. If DpnI will not work, use DNA editing software to choose other restriction endonucleases that cut the template plasmid, but do not cut the PCR product. The combination of using a small amount of template plasmid, endonuclease digestion, and gel extraction minimizes the amount of intact template plasmid that can interfere with downstream recombineering and selection ( Sawitzke et al., 2013 ).

In order to obtain a good gel extraction yield, it is essential to keep the band sharp so that the DNA stays concentrated in a small volume of gel. If the well of an 8-well comb is filled higher than 10 μL, the band can become warped and diffuse. It is better to split the sample across more wells than to overfill a smaller number of wells.

Only run the gel until the desired band is separated enough from any visible side products for excision. Running the gel too long can cause the bands to become warped and diffuse. If the desired band is still too close to easily excise from side products, keep running the gel in 10 minute increments until there is sufficient separation.

Use a blue LED transilluminator while cutting bands out of the gel; do not use an ultraviolet transilluminator, if at all possible.

• Purify the targeting cassette from the gel slice using the Omega Bio-tek Gel Extraction Kit following the manufacturer’s instructions, but elute in 30 μL molecular biology-grade water or 0.2X elution buffer to yield the final targeting cassette. Measure the concentration with a UV-Vis spectrometer and ensure that the concentration is at least 10 ng/μL. Otherwise, repeat steps 1–15. We typically observe yields of 20–50 ng/μL using this protocol.

Eluting with molecular biology-grade water gives the lowest possible salt concentration. However, eluting with unbuffered water could cause lower yields because elution of DNA from silica columns is sensitive to pH. Using 0.2X elution buffer keeps the salt concentration low and also stabilizes the pH to ensure good DNA yields.

SUPPORT PROTOCOL 2

Screening for recombinants by colony pcr.

After selection or counterselection, not all colonies on the plate will be successful recombinants. Many of the colonies will be “false positives” that have only incorporated a truncated portion of the recombineering cassette, that are unmodified parent resulting from a heteromultimer intramolecular recombination ( Fig. 8 ), or that have an inactive, mutant copy of the ccdB gene. Often, when trying to incorporate a particularly large targeting cassette, a large majority of the colonies will be false positives. Colony PCR—in which diagnostic PCR reactions are carried out on individual colonies from the plate—is the quickest and easiest way to identify successful recombinants amongst the false positives. Successful deletion or insertion recombinants can be identified by amplifying the target region with the sequencing primers and looking for an expected size change relative to the umodified parent target region or kan R + ccdB intermediate in the PCR product on an agarose gel. If the expected size change is too small to confidently identify on an agarose gel—generally 200 bp or smaller—successful recombinants can also be identified by amplifying the expected target region with one of the targeting primers and one of the sequencing primers that faces that targeting primer ( Sawitzke et al., 2013 ; Thomason et al., 2014 ). In this case, a PCR product will only be generated if the targeting cassette has been incorporated. Successful recombinants can be identified by evaluating whether there is a band corresponding to the expected PCR product or whether there is no band. For point mutations, the previous two methods will not work, owing to the lack of size change or change in primer binding sites upon recombination. The simplest way to identify successful point mutations is to amplify the target region with the sequencing primers and then Sanger sequence the PCR product ( Sawitzke et al., 2013 ; Thomason et al., 2014 ).

Thermal cycler (e.g., Bio-Rad T100 Thermal Cycler 1861096)

Performing Colony PCR

For recombinants that will have a detectable size change relative to the unmodified parent or kan R +ccdB intermediate, use the sequencing primers to screen the colonies. Otherwise, use one of the targeting primers and the sequencing primer that faces that targeting primer, such that the PCR will only occur if recombination was successful.

This protocol describes how to screen 24 colonies because it is the maximum number of colonies that can be conveniently screened on one agarose gel with two 15-well combs. However, the protocol can be scaled up or down accordingly. Based on our experience, it is advisable to screen at least 8 colonies.

The tubes of LB media serve to store the colonies in an organized manner so that colonies that are identified as successful recombinants can later be retrieved and grown. The colonies can be stored in the LB media in the PCR tubes at 4 °C for at least a week.

• Calculate the annealing temperature for the primers using the New England Biolabs T m Calculator ( https://tmcalculator.neb.com/#!/main ). Ensure the polymerase option is set to “OneTaq Quick-Load 2X Master Mix with Standard Buffer”. Determine the expected size of the PCR product using DNA editing software.
• 94 °C       5 minutes
• 94 °C       15 seconds
• Annealing Temp.   15 seconds
• 68 °C       60 seconds per kb of expected PCR product
• Return to step 2 for 30 cycles
• 68 °C       5 minutes
• 4 °C         hold

Note that the reaction conditions for the OneTaq polymerase mixture are different than those for the Q5 polymerase mixture in Support Protocol 1 . Along with different temperatures and times, the initial denaturation in step 1 of the program is extended to 5 minutes to lyse the bacteria and release template DNA.

Identifying PCR products by gel electrophoresis

• 7. Add 5 μL of GelGreen dye to the molten agarose, swirl the flask to distribute the dye, then pour into a tray in a gel caster with two 15-well combs. Allow the gel to cool and solidify for 15–30 minutes.
• 8. Carefully remove the comb and place the solidified gel into an electrophoresis chamber filled with 0.5X TBE buffer. Ensure that there is sufficient buffer to submerge the entire gel.

The OneTaq Quick-Load 2X Master Mix already contains a green loading dye and a density reagent, so the PCR reaction can just be directly loaded into the well.

• 10. Run the gel at 150 volts for 20 minutes, then visualize on a blue LED transilluminator to search for successful recombinants. If the modification is a point mutation, identify several colonies that have PCR products with the expected size and that have bright bands, purify the PCR products using the Omega Bio-tek Cycle Pure Kit, and submit the purified DNA samples for Sanger sequencing using the sequencing primers and following the service provider’s instructions.
• 11. Once a successful recombinant is identified, either by visually inspecting the colony PCRs on a gel and/or sequencing, return to the PCR tube containing that colony in LB media and grow it up for the next step.

While it is absolutely necessary to sequence the colony PCR products when identifying successful point mutations, it is a good idea to fully sequence other types of modifications once a successful recombinant is identified and chosen by visual inspection of colony PCR products on a gel. Targeting cassettes will sometimes contain random mutations introduced in oligo manufacturing or in PCR reactions that can end up in the final recombinant. It is important to ensure that the final recombinant is free of these unwanted, potentially-deleterious mutations ( Thomason et al., 2014 ).

SUPPORT PROTOCOL 3

Curing the recombineering plasmid.

After a target BAC or strain is modified, the recombineering plasmid is no longer needed and can potentially interfere with downstream experiments if other psc101 plasmids, other tetracycline-resistant plasmids, arabinose induction, or rhamnose induction are used. The psc101 origin of the recombineering plasmid used in these protocols is temperature-sensitive, and can thus be removed or “cured” from the strain by growing at higher temperatures ( H. Wang et al., 2014 ). Alternatively, in the case of target BACs, the mixture of target BAC DNA and recombineering plasmid can be transformed into empty E. coli cells. Colonies from the transformation can then be screened for cells that received the target BAC but not the recombineering plasmid, simply by ensuring that the cells grow in chloramphenicol but not in tetracycline.

LB agar plates with 1X chloramphenicol (see recipe)

42 °C incubator (e.g., VWR Gravity convection incubator 89511–422)

Curing the recombineering plasmid at high temperature

Ensure that the plate does not contain tetracycline. For curing the recombineering plasmid from strains that do not carry BACs, do not use chloramphenicol plates. Instead, look for antibiotics that the strain is resistant to or, less preferably, use plates that have no antibiotic. Strain genotypes that have the letters “gyrA96” are usually resistant to 30 μg/mL nalidixic acid. Strain genotypes that have the letters “rpsL” are usually resistant to 100 μg/mL streptomycin.

• Incubate the plate at 42 °C overnight for ~16 hours or until colonies appear.
• Select any single colony from the plate and streak it onto a new LB agar plate with 1X chloramphenicol.

Occasionally, after the first streak at 42 °C (step 1), the colonies that appear will still contain a few cells that retain the recombineering plasmid. Streaking a second time at 42 °C (step 3) ensures that the resulting colonies are pure and do not contain the recombineering plasmid.

Pick the colony with a sterile pipette tip. Swirl the tip in the 1X chloramphenicol culture first, then use the same tip and swirl it in the 1X tetracycline culture. The tetracycline culture serves as a test to see if the recombineering plasmid is actually cured.

• If the chloramphenicol-containing culture grows but the tetracycline culture does not, thoroughly mix 500 μL of the chloramphenicol culture with 500 μL of sterile 50% glycerol in a sterile 1.7 mL microcentrifuge tube (without a hole in the lid) and store immediately at –80 °C. This glycerol stock is the pure target BAC without the recombineering plasmid. If the tetracycline culture did grow, return to step 5 and test several other colonies.

SUPPORT PROTOCOL 4

Remonomerizing heteromultimers or homomultimers.

Plasmids are prone to forming multimers during recombineering ( Thomason et al., 2007 ). Multimers can interfere with counterselection or result in undesired mixed products. We note that this issue does not occur when targeting the E. coli genome, as E. coli have a system to prevent the formation of multimeric genomes ( Bigot et al., 2007 ). Using the particular protocols described above, we have found that 30–50% of recombinant colonies are heteromultimers resulting from the fusion of unmodified parent BACs and kan R + ccdB intermediate BACs. These heteromultimers are unsuitable for counterselection because they recombine to eliminate the toxic ccdB gene and yield unmodified parent BAC at a frequency much higher than the desired recombineering event ( Fig. 8 ). Furthermore, Basic Protocols 2 and 3 become exceedingly inefficient if the target BAC is a homomultimer because heteromultimeric kan R + ccdB intermediates will then represent nearly 100% of recombinant colonies after the introduction of the kan R + ccdB targeting cassette ( Fig. 9 ). Heteromultimers can be reliably detected by endonuclease digestion, as described in Basic Protocol 1 , steps 34–52. Homomultimers are very difficult to reliably detect with simple methods, as they will appear normal via a PCR- or digestion-based test. Furthermore, it is challenging to ascertain the size of very large, intact plasmids without applying specialized separation techniques, such as pulsed-field gel electrophoresis. The simplest method to detect a homomultimer is to attempt recombineering and then observe that all of the recombinants are heteromultimeric. If heteromultimers or homomultimers are suspected, the issue can be resolved by digesting the BAC with an endonuclease that cuts once, then religating to yield a monomeric species that can be retransformed into DH10B cells ( Thomason et al., 2007 ).

Target BAC or Strain with psc101-gbaA plasmid (supplied by user)

Midiprep DNA isolation kit (e.g., ZymoPURE II Plasmid Midiprep Kit D4200)

Restriction endonucleases (selected by user, e.g., New England Biolabs)

T4 DNA ligase (New England Biolabs M0202S)

New England Biolabs 10-beta Competent E. coli cells (New England Biolabs C3019H)

42 °C water bath (e.g., VWR General Purpose Water Bath 89501–460)

Remonomerizing heteromultimers or homomultimers

• Inoculate a 5 mL culture of LB containing 5 μL of 1000X chloramphenicol stock solution stock solution with the suspected multimer in a sterile 14 mL culture tube. Incubate the culture overnight at 30 °C, shaking at 250 r.p.m. for ~16 hours
• Isolate the target BAC DNA using a midiprep DNA isolation kit, following the manufacturer’s instructions.
• Using DNA editing software, identify a restriction endonuclease that only cuts the target BAC once. Ideally, the endonuclease should leave a >1 bp overhang.
• Digest ~4 μg of the isolated target BAC DNA with the chosen restriction endonuclease, following the manufacturer’s instructions. Ideally, digest the DNA overnight at the appropriate temperature for the enzyme used to ensure that there will be very little uncut multimer remaining in the sample.
• After digestion, remove the restriction endonuclease by purifying the DNA with the Omega Bio-tek Cycle Pure Kit, following the manufacturer’s instructions. Elute the DNA into 30 μL of molecular biology-grade water or into 0.2X elution buffer.

Large vectors can be difficult to religate and transform. Incubating the ligation reaction for extended periods of time helps to maximize the number of religated monomers available for transformation.

• After incubation, heat inactivate the ligation reaction at 65 °C for 10 minutes in a thermal cycler or heat block.

Transforming large vectors can be very difficult and inefficient. We recommend using commercially available high-efficiency competent cells, and transforming them via heat shock. Electroporation would require purifying the ligation to remove salt, which is generally not worth the loss of religated DNA that occurs during purification.

• After recovery, pellet the cells at 10,000 × g for 1 minute. Resuspend the pellet in 50 μL of LB media.
• Streak the resuspended cells on an LB agar plate with 1X chloramphenicol. Incubate the plate at 30 °C overnight for ~16 hours or until colonies appear.
• If the ligase plate has more colonies than the no-ligase control plate, select several colonies and check if they are monomeric or heteromultimeric, as described in Basic Protocol 1 , steps 34–52. If there is no difference between the ligase plate and the no-ligase control plate, redo the protocol.

There is no easy way to distinguish homomultimers from monomers. Therefore, when attempting to remonomerize a suspected homomultimer, select a few colonies from the ligase plate and reattempt recombineering with these colonies in parallel. Remonomerization can be challenging for large BACs >10 kb. Homomultimers can be an issue particularly when making a series of modifications on the same target, and oftentimes it is easier to return to previous recombineering steps and obtain a new target BAC that is monomeric, rather than trying to remonomerize a homomultimer that has formed in the middle of a series of modifications.

REAGENTS AND SOLUTIONS

Lysogeny broth media.

Dissolve 25 g of Dehydrated LB Broth, Miller (e.g., Difco 244620) in 1000 mL of milli-Q water and sterilize by autoclaving. Store at room temperature. Lysogeny broth is sometimes referred to as “Luria-Bertani broth”, “Luria broth”, or just “LB media.”

1000X streptomycin stock solution

Dissolve 1 g of streptomycin sulfate (e.g., MP Biomedicals 100556) in 10 mL of molecular biology-grade water (e.g., Corning 46–000-CM). Sterilize using a 10 mL syringe (e.g., Becton Dickinson 309604) to pass the solution through a 0.2 μm cellulose acetate syringe filter (e.g., VWR 28145–477) into a sterile 15 mL conical tube (e.g., Cellstar 188271). Store at –20 °C.

1000X chloramphenicol stock solution

Dissolve 250 mg of chloramphenicol (e.g., Sigma-Aldrich C0378) in 10 mL of 100% ethanol (e.g., Koptec V1001) in a sterile 15 mL conical tube (e.g., Cellstar 188271). Store at –20 °C.

1000X tetracycline stock solution

Dissolve 100 mg of tetracycline hydrochloride (e.g., Calbiochem 58346) in molecular biology-grade water (e.g., Corning 46–000-CM). Sterilize using a 10 mL syringe (e.g., Becton Dickinson 309604) to pass the solution through a 0.2 μm cellulose acetate syringe filter (e.g., VWR 28145–477) into a sterile 15 mL conical tube (e.g., Cellstar 188271). Protect from light by wrapping the tube in aluminum foil. Store at −20 °C.

1000X kanamycin stock solution

Dissolve 500 mg of kanamycin monosulfate (e.g., Alfa Aesar J61272) in 10 mL of molecular biology-grade water (e.g., Corning 46–000-CM). Sterilize using a 10 mL syringe (e.g., Becton Dickinson 309604) to pass the solution through a 0.2 μm cellulose acetate syringe filter (e.g., VWR 28145–477) into a sterile 15 mL conical tube (e.g., Cellstar 188271). Store at –20 °C.

10% arabinose stock solution

Dissolve 1 g of L-arabinose (e.g., Chem-Impex 01654) in 10 mL of molecular biology-grade water (e.g., Corning 46–000-CM). Sterilize using a 10 mL syringe (e.g., Becton Dickinson 309604) to pass the solution through a 0.2 μm cellulose acetate syringe filter (e.g., VWR 28145–477) into a sterile 15 mL conical tube (e.g., Cellstar 188271). Store at –20 °C.

10% rhamnose stock solution

Dissolve 1 g of L-rhamnose (e.g., MP Biomedicals 102809) in 10 mL of molecular biology-grade water (e.g., Corning 46–000-CM). Sterilize using a 10 mL syringe (e.g., Becton Dickinson 309604) to pass the solution through a 0.2 μm cellulose acetate syringe filter (e.g., VWR 28145–477) into a sterile 15 mL conical tube (e.g., Cellstar 188271). Store at –20 °C.

Super optimal broth with catabolic repression media

Dissolve 3.2 g of Dehydrated SOC broth (e.g., Teknova S0225) in 100 mL of milli-Q water and sterilize by autoclaving. Store at room temperature.

LB agar plates

Dissolve 25 g Dehydrated LB Broth, Miller (e.g., Difco 244620) in 1000 mL of milli-Q water. Next, add 15 g of agar (e.g., Sigma A1296), which will not dissolve until melted in the autoclave. Sterilize by autoclaving, then retrieve the bottle shortly after the autoclave cycle is finished and before the agar solidifies. Allow the bottle to cool with occasional swirling until it is still hot but able to be held by hand for ~15 seconds (~50 °C). Add 1 mL of 1000X antibiotic stock solution(s) and/or 20 mL of 10% arabinose stock solution. Mix the contents by vigorously swirling the bottle, then quickly distribute 15 mL of the molten agarose solution into each 10 cm petri dish (e.g., VWR 25384–094) using a 50 mL serological pipette (e.g., Cellstar 768180) and pipette gun (e.g., Drummond Portable Pipet-Aid XP 4–000-101). Perform the distribution near the base of a Bunsen burner for sterility. Be careful not to introduce bubbles into the plates, as bubbles will make it challenging to reliably identify small bacterial colonies. Allow the plates to solidify at room temperature for ~30 minutes, then return them to the original plastic sleeve and store at 4 °C. Wrap the sleeves of plates in aluminum foil or store them in a dark place if the antibiotic(s) added are light-sensitive.

0.2X elution buffer

Add 2 mL of elution buffer (e.g., Omega Bio-tek PD089; excess buffer is often included with kits) to 8 mL of molecular biology-grade water (e.g., Corning 46–000-CM) in a sterile 15 mL conical tube (e.g., Cellstar 188271). Store at room temperature.

Sterile 50% glycerol

Mix 25 mL of molecular biology-grade water (e.g., Corning 46–000-CM) with 25 mL of glycerol (e.g., Sigma-Aldrich G7893) in a 50 mL conical tube (e.g., Cellstar 227261). Sterilize by autoclaving. Store at room temperature.

3 M sodium acetate

Dissolve 2.5 g of sodium acetate (e.g., Macron 7372) in 10 mL of molecular biology-grade water (e.g., Corning 46–000-CM).

0.5X TBE buffer

Dilute 200 mL of 5X TBE buffer (e.g., Biotium 41006) into 900 mL of milli-Q water. Store at room temperature.

Background Information

The lambda phage red recombination pathway was first characterized using various re combination d eficient lambda phage mutants—hence red αβγ ( Signer, 1971 ). To briefly summarize the proposed functions of these genes during recombineering ( Murphy, 2016 ):

• Red α encodes a 5′→3′ exonuclease, commonly referred to as exo or lambda exonuclease, that degrades one strand of DNA and exposes regions of the formerly double-stranded targeting cassette as single-stranded DNA.
• Red β encodes an annealase—commonly referred to as beta, bet, or β—that coats single-stranded DNA in the targeting cassette and anneals it with homologous single-stranded DNA that is exposed in the target DNA, usually in Okazaki gaps formed during target DNA replication.
• The γ gene encodes a protein, commonly referred to as gam or gamma, that inhibits the E. coli recBCD exonuclease so that it does not destroy both strands of the linear targeting cassette.

While it was known since the 1960s that these genes could mediate general lambda phage recombination, their ability to promote efficient gene replacement using linear PCR cassettes in E. coli was only demonstrated decades later ( Murphy, 1998 ). Shortly thereafter, it was shown that homology arms as short as 42 bp could be used to replace genes using the recET recombination system ( Zhang, Buchholz, Muyrers, & Stewart, 1998 ), which functions similarly to the lambda red system. It was later shown that the more efficient lambda red system could also use short homology arms ( Muyrers, 1999 ). This development was followed by the rapid publication of several related recomb inogenic eng ineering or “recombineering” techniques ( Copeland, Jenkins, & Court, 2001 ; Datsenko & Wanner, 2000 ; Yu et al., 2000 ). Because the lambda red and recET systems can utilize such short homology arms, targeting cassettes can be generated via PCR using 5′ primer overhangs, rather than having to laboriously clone the large, 500–1000 bp homology arms that are necessary for other homologous recombination methods in E. coli ( Hamilton et al., 1989 ; Kong et al., 1999 ; Winans et al., 1985 ). Since the inception of recombineering, many improvements and variations of the method have been developed, including the incorporation of new counterselection schemes, such as ccdA/ccdB ( H. Wang et al., 2014 ), and the development of even faster recombineering methods, such as DIRex ( Näsvall, 2017 ). For a more comprehensive review of the history, proposed mechanisms, and variations of recombineering, see ( Mosberg, Lajoie, & Church, 2010 ; Murphy, 2016 ; Poteete, 2008 ). The protocols described here are based on reports of others ( Datsenko & Wanner, 2000 ; Näsvall, 2017 ; Sawitzke et al., 2013 ; Thomason et al., 2014 ; H. Wang et al., 2014 ; Warming et al., 2005 ), but with modifications that we have found improve the efficiency of the technique and with detailed commentary to assist a novice lab (such as our own when we first began using these approaches) in successfully choosing and applying recombineering methods.

Critical Parameters

While recombineering is quite robust and reliable when performed correctly, there are numerous pitfalls that can hinder success for a new user. In particular, attention must be paid to the quality of targeting cassette DNA, the formation of multimeric plasmids, and the architecture of the target. Many of these parameters are directly addressed in the annotations below the protocols steps.

Targeting cassette DNA quality

If the targeting cassette DNA concentration is too low, it will be difficult to transform sufficient DNA to attain efficient recombineering ( Yu et al., 2000 ). The DNA must also be pure and desalted. Otherwise, the sample can conduct significant current or arc during electroporation and kill the bacterial cells. For DNA cassettes larger than ~2 kb, it is important to remove unwanted DNA truncation products via gel extraction. Shorter truncation products will undergo recombination at a much higher rate than the large desired cassette ( Kuhlman & Cox, 2010 ). Finally, it is essential for the purified targeting cassette DNA to be free of any template plasmid used during the PCR that could cause false positives upon transformation.

Multimerization

Multimers can easily form during recombineering ( Thomason et al., 2007 ), which can then hinder future recombineering steps or applications. One-step recombineering ( Basic Protocol 1 ) does not select against heteromultimers. It is therefore important to verify whether or not the final recombinant is heteromultimeric. In Basic Protocols 2 and 3 , heteromultimeric kan R + ccdB intermediates can form that will generate many false positives during counterselection via intramolecular recombination ( Fig. 8 ). In general, the time and effort required to screen for monomeric intermediates are not justified, as it is easier to simply proceed with three or more colonies with the expectation that at least one will yield successful recombinants. However, if only heteromultimeric kan R + ccdB intermediates form, the parent target BAC is likely a homomultimer ( Fig. 9 ). The homomultimer will either need to be remonomerized ( Support Protocol 4 ), or replaced with a different monomeric clone. It is often easier to find other monomeric clones of the parent when possible, such as when the parent BAC is an intermediate in a series of modifications to be made, than it is to attempt in vitro remonomerization ( Support Protocol 4 ).

Target sequence and repetition

It is critical to have an accurate sequence of the targeting cassette and target BAC/genome so that correct primers and homology arms can be designed. An accurate and complete sequence is also important for identifying any repeats or repetitive regions that can interfere with recombineering in the following ways:

• 1) If one or both of the targeting cassette homology arms is targeted to a region that is repeated in the target, such as commonly used promoters, terminators or polyadenylation signals, a mixture of products can result from the cassette landing in different iterations of the repeat or spanning and thus deleting repeats.
• 2) Repetition in the target can lead to intramolecular recombination that eliminates the ccdB gene without undergoing the desired recombination ( Bird et al., 2011 ).
• 3) Unintentional homology between the middle of the targeting cassette and the target can cause recombination with only a fragment of the targeting cassette, especially because shorter cassettes recombine more efficiently ( Kuhlman & Cox, 2010 ; Lim, Min, & Jung, 2008 ).
• 4) If there are repeated regions within the targeting cassette itself, these regions can undergo recombination and cause deletions or duplications before the cassette is incorporated into the target.

When there is a choice, avoid targeting cassettes with repetition or targeting repetitive regions. Syntenic dot-plots map the regions of homology between two sequences in a 2D plot, and are a useful approach to easily identify repetition and homology ( Noé & Kucherov, 2005 ). To search for problematic repetition, visit https://bioinfo.lifl.fr/yass/index.php (use default settings except E -value threshold = 0.1, window range = 10 to 20000 bp, window incr = 0) to create and examine syntenic dot-plots of:

• 1) The target versus itself: Avoid choosing homology arms within or between repeats. Altenatively, engineer each repeat on separate BACs and assemble the BACs after editing using restriction endonuclease cloning.
• 2) The targeting cassette versus itself: Avoid introducing repetitive elements with the targeting cassette.
• 3) The proposed targeting cassette versus the target: If there is unintended homology, attempt to eliminate or reduce it from the proposed targeting cassette.

When problematic repetition is unavoidable, many colonies will need to be screened during recombineering. Alternatively, traditional methods such as restriction endonuclease cloning will need to be used. Reducing the induction time of the recombineering enzymes has also been shown to reduce unwanted recombination between repeats ( Narayanan, 2008 ).

Troubleshooting

For a summary of troubleshooting tips from throughout this article, see Table 3 .

Troubleshooting table

Understanding Results

With the protocols described here, one can expect a successful recombination frequency of roughly 1 in 10,000–100,000 cells ( Datta et al., 2008 ). Colonies that appear after counterselection are often either successful recombinants or false positives with inactivated ccdB , with the ratio depending on the efficiency of recombineering. With less efficient large cassettes, a larger proportion of colonies will be false positives with inactivated ccdB . We have observed that, with large cassettes, a successful recombinant can generally be identified after screening 8–24 colonies. When recombineering, one can expect to observe dozens to ~100 colonies after selection or after counterselection. Note that it is normal to observe a thin lawn of bacteria around the colonies when higher concentrations of cells are spread on the plate, which is not an issue. If there are hundreds to thousands of colonies on the plate after selection or counterselection, troubleshooting as described in Table 3 is likely required.

Time Considerations

A few days are generally required to plan and prepare for a modification or a series of modifications. Large primers greater than 50 bp sometimes require extra time to manufacture, so they should be ordered well ahead of time. Once a target BAC or target strain is acquired, the targeting cassettes are synthesized, and the psc101-gbaA plasmid and target BAC are transformed together into DH10B cells, recombineering can be performed on a predictable time table. From growing up a colony or glycerol stock of a target BAC+psc101-gbaA strain to identifying successful recombinants on a plate, each iteration of Basic Protocol 1 requires ~3 days excluding any time required for remonomerization, Basic Protocol 2 requires 4 days, and Basic Protocol 3 requires 5 days. Most days involve just a few minutes to inoculate cultures or streak colonies, or ~30 minutes to set up a colony PCR screen that requires 1–2 hours to run in the thermal cycler. However, days that involve electroporation will require one to dedicate 7–8 hours and to adhere to a strict time table, although there will be several growth periods within that time that require less researcher attention.

Significance Statement

Recombineering is a valuable technique for editing DNA plasmids when traditional methods, such as restriction endonuclease cloning or Gibson assembly, are not feasible or provide inadequate precision owing to target DNA size or the lack of unique restriction endonuclease sites. Recombineering is also the premier technique for editing the Escherichia coli genome. Instead of cutting DNA or assembling fragments of DNA in vitro , recombineering uses homologous recombination to replace DNA segments with PCR products in vivo . The technique can enable facile engineering of large DNA or even intact genomes. It can also be a complex and challenging technique to learn, with many pitfalls. Here, we provide simple guidelines for users choosing from the vast menu of recombineering options and introduce the experimental method via three detailed, easy-to-follow protocols.

ACKNOWLEDGEMENT

The authors acknowledge funding from the NIH Director’s New Innovator Award Grant 1DP2GM119162 and NIAMS Grant R01AR071443 (to M.D.S.) and the National Science Foundation Graduate Research Fellowships under Grant No. 1122374 (to L.J.P.).

INTERNET RESOURCES

http://www.biotec.tu-dresden.de/research/stewart/group-page.html

The Sewart Lab website. Source of the psc101-gbaA plasmid and a rich resource for more tips on recombineering and other variations of the recombineering protocol.

https://redrecombineering.ncifcrf.gov/court-lab.html

The Court Lab website. A rich resource for more tips on recombineering and other variations of the recombineering protocol.

https://bioinfo.lifl.fr/yass/yass.php

YASS. A user-friendly online tool for generating and examining syntenic dot-plots. These plots are useful for identifying regions of homology between two sequences in order to spot problematic repeats or unintended homologies.

http://jorgensen.biology.utah.edu/wayned/ape/

The website for downloading “A plasmid Editor” or “ApE”, a free DNA editing software.

https://tmcalculator.neb.com/#!/main

New England Biolabs T m calculator. An online tool from New England Biolabs for calculating primer annealing temperatures for New England Biolabs PCR products.

https://www.thermofisher.com/us/en/home/brands/thermo-scientific/molecular-biology/molecular-biology-learning-center/molecular-biology-resource-library/thermo-scientific-web-tools/multiple-primer-analyzer.html

Multiple Primer Analyzer. An online tool from Thermo Fisher Scientific for predicting the formation of primer dimers, which can inhibit PCR reactions. For difficult PCRs, primers should be redesigned when possible to prevent primer dimer formation.

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Genetic Engineering Essay Guide With 70 Hot Topics

Genetic engineering has been a subject of heated debate. You will find many essays on genetic engineering, asking you to debate for or against, discuss its ethical implications, or emerging congenital disease.

With all these at hand, you may be tempted to opt-out immediately. However, this top-notch guide seeks to make genetics essay writing as fun and as straightforward as possible. Ride along to see the magic!

What Is An Essay on Genetic Engineering?

Now, genetic engineering in itself is the use of biotechnology to manipulate an organism’s genes directly. Therefore, essays on genetics will require students to explore the set of technologies used to change cells’ genetic makeup. These include the transfer of genes within and across species boundaries to produce novel or improved organisms.

We have various areas of genetic engineering, such as:

• Human genetic engineering definition: Deals with genetic engineering techniques applied to humans
• Genetic engineering in plants: Concentrates on genetically modified plant species

Genetic engineering is mostly applied in medicine and thus its technicality. I know this is a field that most students approach with reverence and uttermost humility. Nonetheless, it doesn’t have to be that way. The next few lines might change your opinion on genetic engineering forever!

Why is genetic engineering necessary?

Importance of Genetic Engineering

It is essential in the following ways:

• Ensures that seed companies can protect modified seed varieties as intellectual property.
• Leads to production o organisms with better traits
• Helps maintain the ecosystem

You can see why this field is unavoidable regardless of the negative talk behind it.

Genetically Engineering Plants and Animals – Essay Sample

Young in practice, a little over forty years old, genetic engineering has provided the scientific community with an abundance of knowledge once thought absurd. Genetic engineering means deliberately changing the genome of an organism to acquire some desired traits during its cultivation. On the whole, genetic engineering has a multitude of advantages and disadvantages when it comes to using it on animals and plants; the most prominent advantages include disease resistance, increased crop yields, and a decrease in need for pesticides and antibiotics, whereas disadvantages include the potential for emergence of stronger pathogens, as well as various unexpected consequences. This current paper discusses the pros and cons of using genetic engineering on plants, animals, and provides a synthesis, arguing that, despite its disadvantages, it still serves as a pivotal advantage not only within the scientific community, but also society.

The Advantages of Using Genetic Engineering

The impact of genetic engineering on society can be seen at various aspects, affecting various aspects of social and physical organic life, especially in terms of human beings. The practice consists of the specific selection and removal of genes from organic organisms and inserting them into another. The practice, though still young in practice and not yet deemed completely socially acceptable, makes the possibility of curing diseases once thought incurable a reality, thereby inherently improving the life of both humans and non-human animals. It has many positive effects on society, an example being in Uganda bananas, a main source of caloric intake, are susceptible to the emergence of new diseases that affects their production because of the disease’s potency. Ugandan scientists have successfully used a genetic modification, inserting a pepper gene into bananas, which prevents the fruit from getting the disease (Bohanec, 2015). Furthermore, through genetic engineering, tissue, skin cells, and other forms of organic matter can be grown and used in replacing damaged, worn, or malfunctioning organs and tissues thereby prolonging human life and benefiting their quality of life. The practice helps better advance both the scientific and medical field, both of which are essential in discovering how to better life on Earth.

Genetic engineering, as previously mentioned, can be used to grow and replace damaged tissue or organs, aiding in the betterment and prolonging of human life; it can cure diseases once though incurable, an example being AIDS and cancer. Millions of people around the world suffer from AIDS and cancer, both posing a severe risk to the overall health of the person. More than 900,000 lives were taken by AIDS in 2017 (UNAIDS, 2018). Similarly, over 600,000 were taken by cancer in the following year (NIH, 2018). Genetic engineering makes the possibility of eradicating these diseases a reality. In theory, genetic engineering can help those who suffer from these diseases live longer, healthier, fuller lives by eradicating the disease in its entirety. Though it would not be an easy feat, nor a cheap one, it could still help further advance and better human life and prolong the human life span. People would no longer live in fear of dying from these prolific diseases. Furthermore, genetic engineering, despite the naysayers and opposers of the practice, is another step in organic evolution. From plants to animals, the practice has the chance to achieve strides within scientific history that can greatly benefit the planet in its entirety. From eliminating hunger, to eradicating once prolific diseases, genetic engineering can provide a better, longer, and higher quality of life and tackle bounds once thought impossible the scientific community.

Genetically engineered plants and animals may provide a wide array of benefits that might be pivotal for humanity in the modern world. These benefits include the possibility of developing such plant cultivars that would be resistant to a wide variety of pathogens and diseases caused by microorganisms such as viruses (Ginn, Alexander, Edelstein, Abedi, & Wixon, 2013). If such plant cultivars are created, it might become unnecessary to use chemicals in order to battle these plant diseases. This is clearly a major benefit, since it means better preserving the natural environment and avoiding the use of chemicals that may contaminate soils and waters, as well as kill wildlife.

The Disadvantages of Using Genetic Engineering

The use of genetic engineering to alter plants and animals used in agriculture and husbandry may also have a variety of adverse consequences. For instance, it should be noted that high rates of resistance to disease might have a serious flip side. More specifically, the pathogenic microorganisms (such as bacteria and viruses) can usually mutate quickly in order to adapt to the new conditions. This means that if new cultivars or breeds of plants or animals with high resistance to diseases are created, the pathogens may adapt to these changes in their “hosts” and turn stronger, thus becoming capable of infecting the new cultivars or breeds (Ayres, n.d.). This might again necessitate the use of chemicals or antibiotics; only now stronger drugs or pesticides would be needed. In addition, the old cultivars or breeds may also become infected by the new microorganism strains, and these strains will probably cause more severe diseases in the “original” plants and animals and will be more difficult to cure or prevent.

Another negative possibility is accidentally creating some invasive species that may harm the local ecosystems. For instance, if new plants are made in such a manner that the local species of animals cannot eat them, and then humans lose control over their growth, the new plants may pose a danger to the original plants growing in the given ecosystem, therefore disrupting the ecosystem. For example, in 1984 a patch of seaweed labelled as Caulerpa taxifolia was bred with another robust strain of seaweed identified by scientists as Caulerpa taxifolia (Vahl) C. Agandh . The initial objective was to breed an aquarium plant, however, after a sample escaped in 1984 into the Mediterranean Sea, being found off the coast of both the United States and Australia in 2000, it was found that the strain’s taste was subpar to marine wild life. It was eventually poisoned by the California state government to avoid further damage to marine life and the marine ecosystem and was consequently outlawed by hundreds of countries. The World Conservation Union named it one of the 100 World’s Worst Invasive Alien Species, despite it being manmade (Cellania, 2008).

Finally, there is always the risk of “going too far” when practicing genetic engineering (Bruce & Bruce, 2013). Indeed, it should be noted that the humanity has used various methods of cultivation for millennia in order to breed for specific traits. For example, in 1956, Warwick Kerr, a Brazilian geneticist, imported an aggressive breed of African honeybee to breed with a European species to aid in the decreasing bee population epidemic. Provoked by even the smallest of instigation, after over 26 swarms of the aggressive bee escaped from the apiary in Sao Paulo, they wreaked havoc in North and South America, found in the United States in the early 90s. Nevertheless, genetic engineering is a fast and radical method to change organisms, and very little, if any, data is available to predict the potential adverse impacts of its utilization. It may be difficult to tell when (if at any point) one must stop the process of genetic engineering to avoid unexpected adverse influences of its utilization.

Genetic engineering, despite its disadvantages, can help progress humanity in ways that once seemed impossible. With the environmental and physical epidemics surrounding the planet, the practice can serve as a benefit to resolving the hunger crisis, the preservation of endangered plant and animal species, bringing certain species back from extinction, and so much more. It should be stressed that the utilization of biotechnology and genetic engineering may bring a wide array of significant benefits, which may be of great use to the humanity nowadays. The creation of breeds and cultivars which are immune to disease, resistant to harsh environmental conditions, are cheap to grow, and provide better nutritional value for people might be extremely helpful in reducing the amount of chemicals, pesticides, and antibiotics needed to grow these animals or plants, and, consequently, to help preserve the environment. However, it should also be remembered that genetic engineering might have a wide array of adverse impacts, such as the emergence of new, stronger pathogens, the creation of invasive species, and a multitude of negative consequences that no one knew to expect.

Genetic Engineering Essay Structure

A top-rated genetic engineering essay comes in the manner outlined below:

• Genetic engineering essay introduction: Provide context for your paper by giving a well-researched background on the subject of discussion. Include the thesis statement which will provide the direction of your writing.
• Body: Discuss the main points in detail with relevant examples and evidence from authentic and reliable sources. You can use diagrams or illustrations to support your argument if need be.
• Genetic engineering conclusion: Finalize your paper with a summative statement and a restatement of the thesis statement while showing the genetic engineering process’s implication. Does it add any value to society?

Armed with this great treasure of knowledge, you are good to begin writing your paper. However, we have quality genetic engineering essay topics from expert writers to start you off:

Interesting Genetic Engineering Persuasive Essay Topics

• How human curiosity has led to new advancements and technologies in genetics
• History of genetically modified food
• Discuss the process of genetic engineering in crops
• Evaluate the acceptance of genetically modified crops worldwide
• Analyze the leading countries implementing genetic engineering
• Does genetic engineering produce a desired characteristic?
• What are the legal implications of genetic engineering
• The role of scientists in making the world a better place
• Why coronavirus is a game-changer in the field of genetic engineering
• The effectiveness of genetic engineering as a course in college

Great Topics on the Disadvantages of Genetic Engineering in Humans

• Why changing the sequence of nucleotides of the DNA affects human code structure
• Impact of genetic engineering human lifespans
• Genetic engineering and population control
• Ethical questions to consider in human genetic engineering
• Unintended side effects on humans
• Increasing the risk of allergies
• The foundation of new weapon technologies
• Disadvantages of trait selection before birth
• The greater risk of stillbirth
• Why ladies are at risk with genetic engineering

Why is Genetic Engineering Good Essay Topics

• Genetic engineering and disease prevention
• The creation of a healthy and better society
• Production of drought-resistant crops
• Crop pollen spreads further than expected
• Survival of human species
• Birth of healthy children with desirable traits
• Solving food insecurity problems globally
• Elimination of fertility issues for couples
• Medical advancements as a result of genetic engineering
• Reducing the prevalence of schizophrenia and depression

Good Genetic Engineering Topics

• The development of genetic engineering in the modern world
• Application of ethics in genetic engineering
• Societal class versus genetic engineering
• Impact of genetic engineering on natural selection and adaptation
• Detection of toxins from GMO foods
• Social effects of genetic engineering
• Why people are becoming increasingly resistant to antibiotics
• How gene editing affects the human germline
• Medical treatment opportunities in genetic engineering
• The relationship between molecular cloning and genetic engineering

Impressive Genetic Engineering Research Paper Topics

• Impact of genetic engineering on food supply
• The taste of GMO food versus ordinary food
• GMOs and their need for environmental resources
• Why genetic engineering may face out the use of pesticides
• Reduced cost of living and longer shelf life.
• Growth rates of plants and animals
• Application of genetic engineering on soil bacteria
• New allergens in the food supply
• Production of new toxins
• Enhancement of the environment for toxic fungi

Latest Genetic Engineering Ideas

• The discovery of vaccines through genetic engineering
• Biological warfare on the rise
• Change in herbicide use patterns
• Mutation effects in plants and animals
• Impact of gene therapies
• Does genetic engineering always lead to the desired phenotype?
• Genetic engineering in mass insulin production
• Role of genetic engineering in human growth hormones
• Treating infertility
• Development of monoclonal antibodies

Pro and Cons of Genetic Engineering in Humans Topic Ideas

• Possibility of increased economic inequality
• Increased human suffering
• The emergence of large-scale eugenic programmes
• Rise of totalitarian control over human lives
• The concentration of toxic metals in genetic engineering
• Creation of animal models of human diseases
• Using somatic gene therapy on Parkinson’s disease
• Production of allergens in the food supply
• Redesigning the world through genetic engineering
• Bioterrorism: A study of the issue of emerging infectious diseases

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119 Genetics Research Topics You Must Know About

Put simply, Genetics is the study of genes and hereditary traits in living organisms. Knowledge in this field has gone up over time, and this is proportional to the amount of research.

Right from the DNA structure discovery, a lot more has come out into the open. There are so many genetics research topics to choose from because of the wide scope of research done in recent years.

Genetics is so dear to us since it helps us understand our genes and hereditary traits. In this guide, you will get to understand this subject more and get several topic suggestions that you can consider when looking for interesting genetics topics.

Writing a paper on genetics is quite intriguing nowadays. Remember that because there are so many topics in genetics, choosing the right one is crucial. It will help you cut down on research time and the technicality of selecting content for the topic. Thus, it would matter a lot if you confirmed whether or not the topic you’re choosing has relevant sources in plenty.

What Is Genetics?

Before we even go deeper into genetics topics for research papers, it is essential to have a basic understanding of what the subject entails.

Genetics is a branch of Biology to start with. It is mainly focused on the study of genetic variation, hereditary traits, and genes.

Genetics has relations with several other subjects, including biotechnology, medicine, and agriculture. In Genetics, we study how genes act on the cell and how they’re transmitted from a parent to the offspring. In modern Genetics, the emphasis is more on DNA, which is the chemical substance found in genes. Remember that Genetics cut across animals, insects, and plants – basically any living organism there is.

Tips On How To Write A Decent Research Paper On Genetics

When planning to choose genetics topics, you should also make time and learn how to research. After all, this is the only way you can gather the information that will help you come up with the content for the paper. Here are some tips that can bail you out whenever you feel stuck:

Choosing the topic, nonetheless, is not an easy thing for many students. There are just so many options present, and often, you get spoilt for choice. But note that this is an integral stage/process that you have to complete. Do proper research on the topic and choose the kind of information that you’d like to apply.

Choose a topic that has enough sources academically. Also, choosing interesting topics in genetics is a flex that can help you during the writing process.

On the web, there’s a myriad of information that often can become deceiving. Amateurs try their luck to put together several pieces of information in a bid to try and convince you that they are the authority on the subject. Many students become gullible to such tricks and end up writing poorly in Genetics.

Resist the temptation to look for an easy way of gaining sources/information. You have to take your time and dig up information from credible resources. Otherwise, you’ll look like a clown in front of your professor with laughable Genetics content.

Also, it is quite important that you check when your sources were updated or published. It is preferred and advised that you use recent sources that have gone under satisfactory research and assessment.

Also, add a few words to each on what you’re planning to discuss.Now, here are some of the top genetics paper topics that can provide ideas on what to write about.

Good Ideas For Genetics Topics

Here are some brilliant ideas that you can use as research paper topics in the Genetics field:

• Is the knowledge of Genetics ahead of replication and research?
• What would superman’s genetics be like?
• DNA molecules and 3D printing – How does it work?
• How come people living in mountainous regions can withstand high altitudes?
• How to cross genes in distinct animals.
• Does gene-crossing really help to improve breeds or animals?
• The human body’s biggest intriguing genetic contradictions
• Are we still far away from achieving clones?
• How close are we to fully cloning human beings?
• Can genetics really help scientists to secure various treatments?
• Gene’s regulation – more details on how they can be regulated.
• Genetic engineering and its functioning.
• What are some of the most fascinating facts in the field of Genetics?
• Can you decipher genetic code?
• Cancer vaccines and whether or not they really work.
• Revealing the genetic pathways that control how proteins are made in a bacterial cell.
• How food affects the human body’s response to and connection with certain plants’ and animals’ DNA.

Hot Topics In Genetics

In this list are some of the topics that raise a lot of attention and interest from the masses. Choose the one that you’d be interested in:

• The question of death: Why do men die before women?
• Has human DNA changed since the evolution process?
• How much can DNA really change?
• How much percentage of genes from the father goes to the child?
• Does the mother have a higher percentage of genes transferred to the child?
• Is every person unique in terms of their genes?
• How does genetics make some of us alike?
• Is there a relationship between diets and genetics?
• Does human DNA resemble any other animal’s DNA?
• Sleep and how long you will live on earth: Are they really related?
• Does genetics or a healthy lifestyle dictate how long you’ll live?
• Is genetics the secret to long life on earth?
• How much does genetics affect your life’s quality?
• The question on ageing: Does genetics have a role to play?
• Can one push away certain diseases just by passing a genetic test?
• Is mental illness continuous through genes?
• The relationship between Parkinson’s, Alzheimer’s and the DNA.

Molecular Genetics Topics

Here is a list of topics to help you get a better understanding of Molecular genetics:

• Mutation of genes and constancy.
• What can we learn more about viruses, bacteria, and multicellular organisms?
• A study on molecular genetics: What does it involve?
• The changing of genetics in bacteria.
• What is the elucidation of the chemical nature of a gene?
• Prokaryotes genetics: Why does this take a centre stage in the genetics of microorganisms?
• Cell study: How this complex assessment has progressed.
• What tools can scientists wield in cell study?
• A look into the DNA of viruses.
• What can the COVID-19 virus help us to understand about genetics?
• Examining molecular genetics through chemical properties.
• Examining molecular genetics through physical properties.
• Is there a way you can store genetic information?
• Is there any distinction between molecular levels and subcellular levels?
• Variability and inheritance: What you need to note about living things at the molecular level.
• The research and study on molecular genetics: Key takeaways.
• What scientists can do within the confines of molecular genetics?
• Molecular genetics research and experiments: What you need to know.
• What is molecular genetics, and how can you learn about it?

Human Genetics Research Topics

Human genetics is an interesting field that has in-depth content. Some topics here will jog your brain and invoke curiosity in you. However, if you have difficulty writing a scientific thesis , you can always contact us for help.

• Can you extend your life by up to 100% just by gaining more understanding of the structure of DNA?
• What programming can you do with the help of DNA?
• Production of neurotransmitters and hormones through DNA.
• Is there something that you can change in the human body?
• What is already predetermined in the human body?
• Do genes capture and secure information on someone’s mentality?
• Vaccines and their effect on the DNA.
• What’s the likelihood that a majority of people on earth have similar DNA?
• Breaking of the myostatin gene: What impact does it have on the human body?
• Is obesity passed genetically?
• What are the odds of someone being overweight when the rest of his lineage is obese?
• A better understanding of the relationship between genetics and human metabolism.
• The truths and myths engulfing human metabolism and genetics.
• Genetic tests on sports performance: What you need to know.
• An insight on human genetics.
• Is there any way that you can prevent diseases that are transmitted genetically?
• What are some of the diseases that can be passed from one generation to the next through genetics?
• Genetic tests conducted on a person’s country of origin: Are they really accurate?
• Is it possible to confirm someone’s country of origin just by analyzing their genes?

Current Topics in Genetics

A list to help you choose from all the most relevant topics:

• DNA-altering experiments: How are scientists conducting them?
• How important is it to educate kids about genetics while they’re still in early learning institutions?
• A look into the genetics of men and women: What are the variations?
• Successes and failures in the study of genetics so far.
• What does the future of genetics compare to the current state?
• Are there any TV series or science fiction films that showcase the future of genetics?
• Some of the most famous myths today are about genetics.
• Is there a relationship between genetics and homosexuality?
• Does intelligence pass through generations?
• What impact does genetics hold on human intelligence?
• Do saliva and hair contain any genetic data?
• What impact does genetics have on criminality?
• Is it possible that most criminals inherit the trait through genetics?
• Drug addiction and alcohol use: How close can you relate it to genetics?
• DNA changes in animals, humans, and plants: What is the trigger?
• Can you extend life through medication?
• Are there any available remedies that extend a person’s life genetically?
• Who can study genetics?
• Is genetics only relevant to scientists?
• The current approach to genetics study: How has it changed since ancient times?

Controversial Genetics Topics

Last, but definitely not least, are some controversial topics in genetics. These are topics that have gone through debate and have faced criticism all around. Here are some you can write a research paper about:

• Gene therapy: Some of the ethical issues surrounding it.
• The genetic engineering of animals: What questions have people raised about it?
• The controversy around epigenetics.
• The human evolution process and how it relates to genetics.
• Gene editing and the numerous controversies around it.
• The question on same-sex relations and genetics.
• The use of personal genetic information in tackling forensic cases.
• Gene doping in sports: What you need to know.
• Gene patenting: Is it even possible?
• Should gene testing be compulsory?
• Genetic-based therapies and the cloud of controversy around them.
• The dangers and opportunities that lie in genetic engineering.
• GMOs and their impact on the health and welfare of humans.
• At what stage in the control of human genetics do we stop to be human?
• Food science and GMO.
• The fight against GMOs: Why is it such a hot topic?
• The pros and cons of genetic testing.
• The debates around eugenics and genetics.
• Labelling of foods with GMO: Should it be mandatory?
• What really are the concerns around the use of GMOs?
• The Supreme Court decision on the patent placed on gene discoveries.
• The ethical issues surrounding nurses and genomic healthcare.
• Cloning controversial issues.
• Religion and genetics.
• Behavior learning theories are pegged on genetics.
• Countries’ war on GMOs.
• Studies on genetic disorders.

Get Professional Help Online

Now that we have looked at the best rated topics in genetics, from interesting to controversial topics genetics, you have a clue on what to choose. These titles should serve as an example of what to select.

Nonetheless, if you need help with a thesis, we are available to offer professional and affordable thesis writing services . Our high quality college and university assignment assistance are available to all students online at a cheap rate. Get a sample to check on request and let us give you a hand when you need it most.

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Sustainable Food Technology

Computational modeling for the enhancement of thermosonicated sohphie ( myrica esculenta ) fruit juice quality using artificial neural networks (ann) coupled with a genetic algorithm †.

* Corresponding authors

a Department of Food Engineering & Technology, Central Institute of Technology, Deemed to be University, Kokrajhar, B.T.R., Assam, India E-mail: [email protected] Tel: +91-7002909335

This study investigated the impact of thermosonication on enhancing the nutritional characteristics of juice derived from Sohphie ( Myrica esculenta ) fruits. This investigation introduces an innovative approach utilizing artificial neural networks (ANNs) for the multifaceted optimization of the juice extraction process. Specifically, we focused on determining the most effective extraction parameters for thermosonication, including amplitude (30%, 40%, and 50%), treatment time (15, 30, 45, and 60 min) and temperature (30 °C, 40 °C, and 50 °C). The primary objective of this approach was to augment the nutritional and microbiological properties of Sohphie juice by improving its quality attributes such as ascorbic acid (AA) content, anti-oxidant activity (AOA), total anthocyanin content (TAC), total carotenoid content (TCC), total flavonoid content (TFC), total phenolic content (TPC), total viable count (TVC), and yeast and mould count (YMC). The maximum levels of AA (58.74 ± 3.56 mg/100 mL), AOA (66.11% ± 3.92%), TAC (48.50 ± 4.57 μg mL −1 ), TCC (133.60 ± 5.17 βCE μg mL −1 ), TFC (55.49 ± 3.86 mg quercetin equivalents (QE) per mL), TPC (78.94 ± 4.84 mg gallic acid equivalents (GAE) per mL), TVC (2.44 ± 0.23 log CFU mL −1 ) and YMC (1.01 ± 0.11 log CFU mL −1 ) were obtained in thermosonicated Sohphie juices (TSSJ) under optimal conditions. This study highlights that artificial neural networks (ANNs) coupled with a genetic algorithm (GA) are a beneficial resource for forecasting the extraction efficiency of Sohphie fruit juice (SJ) and suggests that employing thermosonication as a preservation method for SJ can potentially replace traditional thermal pasteurization. This strategy has the potential to reduce or prevent quality deterioration while enhancing the functionality of the juice.

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Computational modeling for the enhancement of thermosonicated Sohphie ( Myrica esculenta ) fruit juice quality using artificial neural networks (ANN) coupled with a genetic algorithm

P. Das, P. K. Nayak and R. K. Kesavan, Sustainable Food Technol. , 2024, Advance Article , DOI: 10.1039/D4FB00004H

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Should We Change Species to Save Them?

When traditional conservation fails, science is using “assisted evolution” to give vulnerable wildlife a chance.

Credit... Photo illustration by Lauren Peters-Collaer

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By Emily Anthes

Photographs by Chang W. Lee

This story is part of a series on wildlife conservation in Australia, which Emily Anthes reported from New York and Australia, with Chang W. Lee.

• Published April 14, 2024 Updated April 16, 2024

For tens of millions of years, Australia has been a playground for evolution, and the land Down Under lays claim to some of the most remarkable creatures on Earth.

It is the birthplace of songbirds, the land of egg-laying mammals and the world capital of pouch-bearing marsupials, a group that encompasses far more than just koalas and kangaroos. (Behold the bilby and the bettong!) Nearly half of the continent’s birds and roughly 90 percent of its mammals, reptiles and frogs are found nowhere else on the planet.

Australia has also become a case study in what happens when people push biodiversity to the brink. Habitat degradation, invasive species, infectious diseases and climate change have put many native animals in jeopardy and given Australia one of the worst rates of species loss in the world.

In some cases, scientists say, the threats are so intractable that the only way to protect Australia’s unique animals is to change them. Using a variety of techniques, including crossbreeding and gene editing, scientists are altering the genomes of vulnerable animals, hoping to arm them with the traits they need to survive.

“We’re looking at how we can assist evolution,” said Anthony Waddle, a conservation biologist at Macquarie University in Sydney.

It is an audacious concept, one that challenges a fundamental conservation impulse to preserve wild creatures as they are. But in this human-dominated age — in which Australia is simply at the leading edge of a global biodiversity crisis — the traditional conservation playbook may no longer be enough, some scientists said.

“We’re searching for solutions in an altered world,” said Dan Harley, a senior ecologist at Zoos Victoria. “We need to take risks. We need to be bolder.”

The extinction vortex

The helmeted honeyeater is a bird that demands to be noticed, with a patch of electric-yellow feathers on its forehead and a habit of squawking loudly as it zips through the dense swamp forests of the state of Victoria. But over the last few centuries, humans and wildfires damaged or destroyed these forests, and by 1989, just 50 helmeted honeyeaters remained, clinging to a tiny sliver of swamp at the Yellingbo Nature Conservation Reserve.

Intensive local conservation efforts, including a captive breeding program at Healesville Sanctuary, a Zoos Victoria park, helped the birds hang on. But there was very little genetic diversity among the remaining birds — a problem common in endangered animal populations — and breeding inevitably meant inbreeding. “They have very few options for making good mating decisions,” said Paul Sunnucks, a wildlife geneticist at Monash University in Melbourne.

In any small, closed breeding pool, harmful genetic mutations can build up over time, damaging animals’ health and reproductive success, and inbreeding exacerbates the problem. The helmeted honeyeater was an especially extreme case. The most inbred birds left one-tenth as many offspring as the least inbred ones, and the females had life spans that were half as long, Dr. Sunnucks and his colleagues found.

Without some kind of intervention, the helmeted honeyeater could be pulled into an “extinction vortex,” said Alexandra Pavlova, an evolutionary ecologist at Monash. “It became clear that something new needs to be done.”

A decade ago, Dr. Pavlova, Dr. Sunnucks and several other experts suggested an intervention known as genetic rescue , proposing to add some Gippsland yellow-tufted honeyeaters and their fresh DNA to the breeding pool.

The helmeted and Gippsland honeyeaters are members of the same species, but they are genetically distinct subspecies that have been evolving away from each other for roughly the last 56,000 years. The Gippsland birds live in drier, more open forests and are missing the pronounced feather crown that gives helmeted honeyeaters their name.

Genetic rescue was not a novel idea. In one widely cited success, scientists revived the tiny, inbred panther population of Florida by importing wild panthers from a separate population from Texas.

But the approach violates the traditional conservation tenet that unique biological populations are sacrosanct, to be kept separate and genetically pure. “It really is a paradigm shift,” said Sarah Fitzpatrick, an evolutionary ecologist at Michigan State University who found that genetic rescue is underused in the United States.

Crossing the two types of honeyeaters risked muddying what made each subspecies unique and creating hybrids that were not well suited for either niche. Moving animals between populations can also spread disease, create new invasive populations or destabilize ecosystems in unpredictable ways.

Genetic rescue is also a form of active human meddling that violates what some scholars refer to as conservation’s “ ethos of restraint ” and has sometimes been critiqued as a form of playing God.

“There was a lot of angst among government agencies around doing it,” said Andrew Weeks, an ecological geneticist at the University of Melbourne who began a genetic rescue of the endangered mountain pygmy possum in 2010. “It was only really the idea that the population was about to go extinct that I guess gave government agencies the nudge.”

Dr. Sunnucks and his colleagues made the same calculation, arguing that the risks associated with genetic rescue were small — before the birds’ habitats were carved up and degraded, the two subspecies did occasionally interbreed in the wild — and paled in comparison with the risks of doing nothing.

And so, since 2017, Gippsland birds have been part of the helmeted honeyeater breeding program at Healesville Sanctuary. In captivity there have been real benefits, with many mixed pairs producing more independent chicks per nest than pairs composed of two helmeted honeyeaters. Dozens of hybrid honeyeaters have now been released into the wild. They seem to be faring well, but it is too soon to say whether they have a fitness advantage.

Monash and Zoos Victoria experts are also working on the genetic rescue of other species, including the critically endangered Leadbeater’s possum, a tiny, tree-dwelling marsupial known as the forest fairy. The lowland population of the possum shares the Yellingbo swamps with the helmeted honeyeater; in 2023, just 34 lowland possums remained . The first genetic rescue joey was born at Healesville Sanctuary last month.

The scientists hope that boosting genetic diversity will make these populations more resilient in the face of whatever unknown dangers might arise, increasing the odds that some individuals possess the traits needed to survive. “Genetic diversity is your blueprint for how you contend with the future,” Dr. Harley of Zoos Victoria said.

Targeting threats

For the northern quoll, a small marsupial predator, the existential threat arrived nearly a century ago, when the invasive, poisonous cane toad landed in eastern Australia. Since then, the toxic toads have marched steadily westward — and wiped out entire populations of quolls, which eat the alien amphibians.

But some of the surviving quoll populations in eastern Australia seem to have evolved a distaste for toads . When scientists crossed toad-averse quolls with toad-naive quolls, the hybrid offspring also turned up their tiny pink noses at the toxic amphibians.

What if scientists moved some toad-avoidant quolls to the west, allowing them to spread their discriminating genes before the cane toads arrived? “You’re essentially using natural selection and evolution to achieve your goals, which means that the problem gets solved quite thoroughly and permanently,” said Ben Phillips, a population biologist at Curtin University in Perth who led the research.

A field test, however, demonstrated how unpredictable nature can be. In 2017, Dr. Phillips and his colleagues released a mixed population of northern quolls on a tiny, toad-infested island. Some quolls did interbreed , and there was preliminary evidence of natural selection for “toad-smart” genes.

But the population was not yet fully adapted to toads, and some quolls ate the amphibians and died, Dr. Phillips said. A large wildfire also broke out on the island. Then, a cyclone hit. “ All of these things conspired to send our experimental population extinct,” Dr. Phillips said. The scientists did not have enough funding to try again, but “all the science lined up,” he added.

Advancing science could make future efforts even more targeted. In 2015, for instance, scientists created more heat-resistant coral by crossbreeding colonies from different latitudes . In a proof-of-concept study from 2020, researchers used the gene-editing tool known as CRISPR to directly alter a gene involved in heat tolerance.

CRISPR will not be a practical, real-world solution anytime soon, said Line Bay, a biologist at the Australian Institute of Marine Science who was an author of both studies. “Understanding the benefits and risks is really complex,” she said. “And this idea of meddling with nature is quite confronting to people.”

But there is growing interest in the biotechnological approach. Dr. Waddle hopes to use the tools of synthetic biology, including CRISPR, to engineer frogs that are resistant to the chytrid fungus, which causes a fatal disease that has already contributed to the extinction of at least 90 amphibian species.

The fungus is so difficult to eradicate that some vulnerable species can no longer live in the wild. “So either they live in glass boxes forever,” Dr. Waddle said, “or we come up with solutions where we can get them back in nature and thriving.”

Unintended consequences

Still, no matter how sophisticated the technology becomes, organisms and ecosystems will remain complex. Genetic interventions are “likely to have some unintended impacts,” said Tiffany Kosch, a conservation geneticist at the University of Melbourne who is also hoping to create chytrid-resistant frogs . A genetic variant that helps frogs survive chytrid might make them more susceptible to another health problem , she said.

There are plenty of cautionary tales, efforts to re-engineer nature that have backfired spectacularly. The toxic cane toads, in fact, were set loose in Australia deliberately, in what would turn out to be a deeply misguided attempt to control pest beetles.

But some environmental groups and experts are uneasy about genetic approaches for other reasons, too. “Focusing on intensive intervention in specific species can be a distraction,” said Cam Walker, a spokesman for Friends of the Earth Australia. Staving off the extinction crisis will require broader, landscape-level solutions such as halting habitat loss, he said.

Moreover, animals are autonomous beings, and any intervention into their lives or genomes must have “a very strong ethical and moral justification” — a bar that even many traditional conservation projects do not clear, said Adam Cardilini, an environmental scientist at Deakin University in Victoria.

Chris Lean, a philosopher of biology at Macquarie University, said he believed in the fundamental conservation goal of “preserving the world as it is for its heritage value, for its ability to tell the story of life on Earth.” Still, he said he supported the cautious, limited use of new genomic tools, which may require us to reconsider some longstanding environmental values.

In some ways, assisted evolution is an argument — or, perhaps, an acknowledgment — that there is no stepping back, no future in which humans do not profoundly shape the lives and fates of wild creatures.

To Dr. Harley, it has become clear that preventing more extinctions will require human intervention, innovation and effort. “Let’s lean into that, not be daunted by it,” he said. “My view is that 50 years from now, biologists and wildlife managers will look back at us and say, ‘Why didn’t they take the steps and the opportunities when they had the chance?’”

Emily Anthes is a science reporter, writing primarily about animal health and science. She also covered the coronavirus pandemic. More about Emily Anthes

Chang W. Lee has been a photographer for The Times for 30 years, covering events throughout the world. He is currently based in Seoul. Follow him on Instagram @nytchangster . More about Chang W. Lee

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The Ethical Issues of Genetic Engineering

Introduction, historical context, political context, social context, ethical evaluation, ethical questions, reference list.

Genetic engineering is a scientific achievement that has led to the development of new ethical issues. Genetic engineering has been a subject of controversy because a lot of people are not comfortable with the technology.

The ethical issues are more evident when it comes to cases of genetic engineering on the human tissue. Historically, the process has been conducted in the west. It has become easier to conduct genetic engineering in plants, animals, and humans due to developments in science.

A case study to consider in the relationship between science and ethics is Argentina. The government allowed several manufactures of genetically modified (GM) seeds to operate within the country due to increasing debts.

The manufacturers were given permits and the produced seeds were supplied to farmers for free. The seeds were of a wide variety, including maize, soya, and sunflower. GM soya seeds became common and the country was able to export its soya produce within a short while (Burachik, 2012).

Thus, the government was able to gain through this strategy. Despite this, ethical questions arose about whether the decision made could be considered moral or not.

The ethical questions arising from science are based on two concepts. The first concept is whether science is a danger in itself. The knowledge that arises from science can be a risk (Griffiths & Stotz, 2013). Secondly, an ethical issue arises based on what the long-term effects of science might be.

The idea of improving nature is considered to be a dangerous choice. Thus, it is unethical to change nature. It is easier to establish the ethical argument by raising an extrinsic question that is related to the long-term effects of GM crops makes.

Thus, such a question will be able to inform whether a choice can be considered to be ethical or not. The consequences that arise from the decision are also looked at in detail. Different results may be achieved. Determining whether the choice taken is ethical depends on weighing both options.

An option that has more positive consequences is always considered to be ethical and ideal to choose. In Argentina’s case, the ethical nature of its actions was defined by the financial costs involved. The country was to gain more from the decision to grow GM crops.

For a long time, science was not considered to be a concept that could be tied to ethical considerations. This changed and the social, political, individual, and practical effects were discussed in many forums dealing with the philosophy of science.

Genetic engineering is a science that has the highest potential of changing human lives. Historically, genetic engineering has led to the development of new ethical arguments because GM crops have varied implications that can affect a country as a whole.

In Argentina, the government was able to increase its imports and employ more people in the agricultural sector (Burachik, 2012).

Scientific research has always enjoyed independence when it comes to the expected results. Thus, scientists could conduct any experiment they wanted as long as they were not limited by funds. It is during the 1980s that it was realized that scientific research should be restricted.

The restriction also considered how science should be limited and within what limits (Light & De-Shalit, 2003). It is easier to know the consequences of genetic engineering through rational means. Initially, genetic engineering was witnessed within the field of agriculture.

It was conducted to increase food production by producing better crops that could survive harsh weather conditions. Later, it also involved human genetic engineering. Thus, there was a need to consider the ethical implications of genetic engineering (Stock & Campbell, 2000).

Genetically modified crops always raise political issues. The debate is hotter where the crops are made for human consumption. A political issue arises on whether to let the crops into the country or not.

Many people have questioned the health risks that arise from genetically modified crops, thus it is the politicians who have to ensure that the interests of the people are met and their safety is assured (Haugen, 2013).

GM crops are usually cheaper and have high yields when planted. This is advantageous because it is an economic advantage to a country and its citizens.

Various negative issues arise, despite the advantages of GM crops because the growth of GM crops intensifies pressure on unspoiled nature areas such as forests and grasslands.

GM crops tend to easily adapt into many environmental conditions, thus large tracts of land are set to maximize on the benefits (Burachik, 2012). The growth of GM crops affects various political aims within a country. In many countries, nature conservation is the duty of the government.

In Argentina’s case, the growth of soya became a political issue due to the land that was required for its growth. Its growth spread so rapidly that more than 14 million hectares of land were covered by the crop within two years.

The government established policies that allowed for the eviction of people from the land that was considered suitable for agricultural production after the establishment of the nation state of Argentina in 1853.

Moreover, an economic model was also adopted to encourage exportation and acquisition of foreign aid. The government was also involved in the acquisition of permits to plant GM crops comprising of soya, cotton, sunflower, potatoes, maize, and wheat.

Neither the public nor the Congress was informed about this decision. Thus, it can be seen that political problems would have emerged if this policy was considered by Congress or the public.

Moreover, the government also considered the ethical issues that would have come up due to this policy. Thus, they chose not to divulge the information about the permits (Burachik, 2012).

The commission set by the government to consider biotechnology was comprised of representatives from biotechnology companies. This scenario would not come up with appropriate ethical considerations because most of representatives wanted growth of GM crops just for personal profits.

Political implications always arise due to GM foods. Such crops can have negative implications within a state. In Argentina’s case, the crops began to take larger tracts of land.

There was a risk of social justice being compromised because the government did not care about the implications of the GM crops. Exports from soya were sufficient to pay back its debts, thus the government saw no need to establish better policies to control the growth of GM crops.

The citizens also gained due to this decision, thus it was in the best interest of the country. On the other hand, an individualized contract-based ethics arises whereby the production of GM crops is against nature.

Thus, the government should not be involved in the production of GM crops because they interfere with nature (Laurie, 2002).

GM crops usually tend to use methods that pollute the environment. Growth of GM crops involves use of advanced agricultural practices. In Argentina’s case, farmers were given both seeds and fertilizers to grow the crops.

These fertilizers had health risks and they polluted the environment in the long run. Moreover, less efficient eco-farming strategies were promoted. The methods used for agricultural production used various methods that facilitated increased productivity.

Conservation of biodiversity also became an issue. GM plants have an accelerated growth rate, thus they can encroach on a large piece of land within a short time.

The fertilizers and chemicals used may also affect the surrounding plants and animals. For instance, ploughed grasslands can lead to loss of important biodiversity. The other risks involved were theoretical in nature.

The government’s decision can be seen as unethical if questions are raised about the potential risks GM crops have on humans. Information in this regard could only be obtained through empirical means.

Experimentation and experience were the best means to establish this information (Barry, 2011). Cultivation of genetic crops also leads to spread of genetic engineering. This becomes an ethical issue for countries that have not legalized the importation or sale of GM crops.

Such fears are usually faced by government agencies dealing with rural development. GM crops require modern methods of agricultural production, thus people in rural areas will lose their source of income if GM crops are promoted.

The social impacts of GM foods are always considered before permits are given to develop the foods in most western countries. Other food crops can also be affected through jumping genes and pollen flight. This can lead to disastrous consequences, such a limiting food production in the future.

Thus, a democratic decision should be reached through public debate about the implications of GM crops. Establishing a green genetic engineering strategy would be an effective step to begin with (Derr & McNamara, 2003).

The ethical implications within the society arise based on how people will be affected. In Argentina, the government’s decisions can be considered as illegal, but they were ethical to an extent. The government’s decisions, though not revealed to the public, were for the greater good of the public.

Socially, there were gains and losses expected. GM crops are used at the expense of natural crops. Intensive research is usually done to come up with GM crops.

Thus, natural plants will lose their role in life if GM foods. It is a societal obligation to preserve nature. If GM crops are allowed to flourish, then the society will lose its role in protecting nature (Bennett et al., 2013).

Philosophers in the western world have been interested in the development and systemization of the sciences in relation to genetic engineering. There are two general thoughts that have been used to explain how the actions are viewed. These are the utilitarian and Aristotelian thoughts.

Aristotelian uses the belief of good reason to bring out the forces that influence the direction of the actions. Good reasons are always given to explain the reason behind an action, or an event (Light & De-Shalit, 2003).

An ultimate goal is always pursued, thus less credit is given to the negative effects of the action. Such a scenario can be seen with genetic engineering. The larger picture shows that genetic engineering has negative consequences.

Thus, genetic engineers try to show that the process is beneficial and done with good intentions. The goals already achieved through genetic engineering have been helpful to the human race. It is for this reason that genetic engineering has grown and evolved over the years.

Many people ignore the greater consequences of the process. It is as a result of this realization that it becomes important to consider the ethical implications of genetic engineering.

There is always an evaluation of the reasons explaining what genetic engineering seeks to achieve and the product of the process (Reiss & Straughan, 2001).

One the other hand, utilitarian beliefs do not consider the actions of an individual as resulting from either good or bad decisions, but only with a maximization of the agent’s abilities.

This can be applied to genetic engineering where one can view genetic engineering as using knowledge to its maximum. In Argentina’s case, the actions were not specifically for good or bad reasons.

The activities were conducted to ensure that maximum gains were achieved from the knowledge of genetic engineering. Thus, ethical concerns on GM crops arise depending on the implications of GM crops and not the use genetic engineering.

A closer analysis of the field of science would reveal that people always depend on their practical knowledge. This is then utilized when making a judgment on whether something is good or bad.

Ethical considerations are sometimes based on established norms within the society (Frey & Wellman, 2007). Norms are able to describe what rules are applicable within different contexts. The ethical considerations arising from genetic engineering relate to norms within the society.

It determines how certain beliefs are upheld at the expense of other beliefs. It is hard to accept genetic engineering as ethical if the basis of the science is irrational.

The goals of science can be equated to the goals of life. For both concepts, the end involves improvement of human life (Reiss & Straughan, 2001).

Problems are bound to arise more often in cases of cash crops that grow at the expense of food crops. Genetic modification is allowed on cash crops because of their economic importance.

Scientists usually view ethics as essential to their practice and identity. Despite this, their ethical beliefs can change according to current conditions in society.

Thus, an ethical risk can arise from GM crops whereby it could lead to increased research on genetic engineering on humans (Mizzoni, 2010). Ethical questions also arise on whether it is necessary to genetically modify crops. Naturally, such crops can grow in some environments.

The use of genetic engineering makes the process cheaper because crops are made adaptable to different environments and to yield better products. Though it is cheaper, the negative consequences of this decision can be realized in future.

In the case study, new types of pests have appeared because of the genetically modified crops. Initially, it was thought that such an attack would not occur. This only proves that GM crops are not always the best option (Burachik, 2012).

Many of the ethical and moral debates have followed a one-dimensional strategy whereby they are concerned with a single and a specific application of genetic engineering. Human application of this technology has been given significant coverage in comparison to GM crops.

Research on the implications of genetic engineering on animals, plants, and microorganisms has been largely overlooked. If GM crops are encouraged, then the future will be bleak where most food, animals, or humans will be genetically modified (Nordgren, 2001).

Moral and ethical concerns are effective in controlling public opinion. The public will not easily support an idea if it is considered immoral. Thus, concerns have developed that various biotechnology techniques would fail if not given public acceptance.

Philosophy has been used in the explanation of nature and how to interact with it. An important example is the stoic philosophy that describes that humans have to live with nature as it is (Mizzoni, 2010). It is morally wrong for humans to interfere with nature for their own benefit.

Genetic engineering is seen as the most effective way to interfere with nature because genetic materials are the basic structures that comprise humans, plants, and animals.

The human body and its parts can be seen as a system that works in unison. The different parts play different roles to establish a balance in the human body. The same can be said about nature. Each aspect of nature has its own role to play.

Thus, a balance is established to facilitate the survival of man and his dependence on nature. If nature were to be reconfigured through genetic engineering, then there would be a loss of this balance.

For instance, genetic modification in humans can result in the production of a superhuman. If such a human procreates, then it would lead to a situation where more people have genetically engineered genes resulting from his offspring (Yashon & Cummings, 2012).

Thus, a problem may exist within the individual’s genetic pool and researchers are not aware. The same can be said about GM crops. Their use may result in negative consequences as the case was in Argentina whereby new strains of pests emerged.

A survey conducted in the UK to determine public opinion about GM crops found that 70 percent of the total respondents considered it morally wrong. Thus, globally, the beliefs on genetic engineering depend on individual values. People tend to believe that biotechnology is wrong.

In some cases, this is attributed to lack of knowledge of how genetic modification is done. For most people, they consider the issues that can arise from GM crops to be the same with genetic modification of humans (Haugen, 2013).

The decision to depend on ethics may have negative consequences as well. Something may be considered unethical, but it can lead to improvements.

In conclusion, genetic engineering is a scientific breakthrough that has led to developments in biotechnology. Growth and consumption of GM crops have been on the increase, despite little regard for the consequences.

Thus, ethical issues arise as people try to determine whether GM crops are good or bad for humans. Genetic engineering can have very many dangers, but such fears will reduce once it is realized that everything has the potential to be harmful.

Thus, the issues arising due to GM crops can be related to the ethical issues resulting from science.

Barry, VE 2011, Bioethics: At the beginning and end of life, Wadsworth, Belmont, CA.

Bennett, AB, Chi-Ham, C, Barrows, G, Sexton, S, & Zilberman, D, 2013, ‘Agricultural biotechnology: economics, environment, ethics, and the future,’ Annual Review of Environment & Resources, vol. 38, no. 1, pp. 249-279.

Burachik, M 2012, ‘Regulation of GM crops in Argentina’, GM Crops & Food, vol. 3, no.1, pp. 48-51.

Derr, PG. & McNamara, EM 2003, Case studies in environmental ethics, Rowman & Littlefield, Lanham, MD.

Frey, RG, & Wellman, CH 2007, A companion to applied ethics, John Wiley & Sons, Oxford.

Griffiths, P, & Stotz, K 2013. Genetics and philosophy : an introduction, Cambridge University Press, New York, NY.

Haugen, H M 2013, ‘Human rights in natural science and technology professions’ codes of ethics?’, Business & Professional Ethics Journal, vol. 32, no. 1/2, pp. 49-76.

Laurie, GT 2002, Genetic privacy : a challenge to medico-legal norms, Cambridge University Press, New York, NY.

Light, A, & De-Shalit, A, 2003, Moral and political reasoning in environmental practice, MIT, Cambridge, MA

Mizzoni, J 2010, Ethics: the basics, Wiley-Blackwell, West Sussex, UK.

Nordgren, A, 2001, Responsible genetics: the moral responsibility of geneticists for the consequences of human genetics research, Kluwer Academic Publishers, Boston, MA

Reiss, MJ, & Straughan, R 2001, Improving nature?: the science and ethics of genetic engineering, Cambridge University Press, New York, NY.

Stock, G, & Campbell, JH, 2000, Engineering the human germline: an exploration of the science and ethics of altering the genes we pass to our children, Oxford University Press, New York, NY.

Yashon, RK, & Cummings, MR 2012, Human genetics and society, 2nd ed, Brooks/Cole, Belmont, MA.

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IvyPanda. (2023, November 22). The Ethical Issues of Genetic Engineering. https://ivypanda.com/essays/genetic-engineering-2/

"The Ethical Issues of Genetic Engineering." IvyPanda , 22 Nov. 2023, ivypanda.com/essays/genetic-engineering-2/.

IvyPanda . (2023) 'The Ethical Issues of Genetic Engineering'. 22 November.

IvyPanda . 2023. "The Ethical Issues of Genetic Engineering." November 22, 2023. https://ivypanda.com/essays/genetic-engineering-2/.

1. IvyPanda . "The Ethical Issues of Genetic Engineering." November 22, 2023. https://ivypanda.com/essays/genetic-engineering-2/.

Bibliography

IvyPanda . "The Ethical Issues of Genetic Engineering." November 22, 2023. https://ivypanda.com/essays/genetic-engineering-2/.

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Two students named Goldwater Scholars

• Student Awards
• Weinberg College

Two undergraduate researchers from Northwestern University have earned the 2024 Barry Goldwater Scholarship, an honor that supports students who intend to pursue careers in the natural sciences, mathematics and engineering.

As Goldwater Scholars, Kate Carver and Melany Morales represent the scientific talent essential to ensuring the U.S. maintains its global competitiveness and security, according to the Department of Defense (DOD) National Defense Education Programs. The DOD partners with the Goldwater Foundation on the scholarship program.

This year, a total of 438 Goldwater Scholars were chosen from 1,353 science, engineering and mathematics students nominated by 446 colleges and universities across the nation. Many of the scholars have published their research in leading professional journals and have presented their work at professional society conferences, and almost all plan to obtain a Ph.D.

“Goldwater winners use their experience as a pathway into top graduate schools and fellowships in the U.S. and abroad,” said LaTanya Veronica Williams, associate director for STEM in the Office of Fellowships and the campus representative for the program. “Kate and Melany are stellar students with high levels of resilience and maturity. I am sure that they will follow in the path of previous Goldwater winners and go forward to make great contributions to their fields of study.”

Melany Morales

Morales grew up in a bilingual household in Caracas, Venezuela, and often spent time in her mom’s elementary school classroom, watching how the students learned foundational skills. These experiences sparked Morales’ interest in developmental psychology, especially in language and how social interactions influence learning during development.

Now, Morales, a third-year neuroscience and psychology major, works in three labs, including Northwestern’s Infant and Child Development Center and the Language, Education and Reading Neuroscience (LEARN) lab, where she is working on her honors thesis, as well as Northeastern University’s PINE Lab.

Looking ahead, she hopes to continue to research how early childhood experiences and neuroplasticity shape emerging language and social-emotional skills. 

“This scholarship signifies not only personal achievement, but also the supportive community that has propelled me forward,” Morales said. “I am profoundly grateful for the opportunities it presents and the relationships it has fostered.”

Kate Carver

Carver learned the power of genomic medicine at a young age. Soon after her younger sister, Ella, was born, her family realized that Ella had a developmental delay, but doctors were unable to pinpoint what caused it.

Only when the family was able to have a special genetic sequencing done did they learn that variants on a gene important for neural development were responsible for Ella’s developmental delay. The experience led Carver to pursue a path to help genetics empower other families, too.

For the past two years, Carver, a third-year neuroscience major with a data science minor, has worked with the Perera Lab at Northwestern University Feinberg School of Medicine. There, she studies how patients’ genetics can be used to predict how they will respond to drugs, with the goal of improving treatments and patient outcomes.

“What keeps me going is knowing that there's a little bit of magic in the work we're doing, that it has a significant impact on families downstream, on patients like Ella,” Carver said.

Learn more about the Goldwater Scholarship and other opportunities by contacting Northwestern’s  Office of Fellowships .

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Image caption: Clockwise from left: Shubhayu Bhattacharyay, Min Jae Kim, James Occean, and Michael Xie

Two Johns Hopkins alumni, two graduate students named Paul and Daisy Soros Fellows

They are among 30 recipients of the fellowships, which honor immigrants and children of immigrants with exceptional potential to make a difference in their fields.

By Aleyna Rentz

Four Johns Hopkins affiliates have been awarded the Paul and Daisy Soros Fellowships for New Americans . One of the most competitive scholarships in the United States, the Soros Fellowship honors the contributions of immigrants and children of immigrants to the United States. This year, 30 fellows were chosen from over 2,300 applicants. Fellows are awarded up $90,000 in financial support and are chosen for their potential to make significant contributions to their academic field.

This year's awardees from Johns Hopkins are Shubhayu Bhattacharyay, Engr '20; Min Jae Kim, Engr '22; James Occean, who is pursuing a master's degree in bioinformatics; and Michael E. Xie, who is pursuing an MD/PhD in the Medical Scientist Training Program.

Shubhayu Bhattacharyay, Engr '20

Shubhayu Bhattacharyay was born in Kolkata, India, and spent his early childhood in Thailand and Vietnam before settling in the South Bay of Los Angeles. At Johns Hopkins University, Bhattacharyay double majored in biomedical engineering and applied mathematics and statistics with a minor in Spanish. He was supported by the Milken Scholars Program and graduated with full departmental and Tau Beta Pi honors. As an undergraduate, Bhattacharyay founded Auditus Technologies, a company inventing individualizable, accessible hearing devices for adults living with dementia.

Bhattacharyay started to consider a medical career in the summer after his first year at Hopkins, when he met traumatic brain injury (TBI) survivors participating in a brain-computer interface study. Their stories motivated Bhattacharyay to think of ways his interest in computational neuroscience might contribute towards an improved quality of life after TBI. Mentored by Robert Stevens , director of the Johns Hopkins Division of Informatics, Integration, and Innovation and an associate professor of anesthesiology and critical care medicine, Bhattacharyay invented and published results from the first computational bedside system to sense and classify motor function in TBI patients in the intensive care unit.

In 2020, Bhattacharyay received a Gates Cambridge Scholarship to pursue a PhD in clinical neurosciences at the University of Cambridge under the supervision of professors Ari Ercole and David Menon. For his thesis, Bhattacharyay developed artificial intelligence methods which improve the detail of information provided for prognostic counseling and suggest individually optimized treatment plans during the ICU management of TBI. His work has generated publications in leading digital health and neurotrauma journals, open access software packages, and invited talks at international conferences. During his graduate studies, Bhattacharyay volunteered at Headway Cambridge and Peterborough, a charity-run rehabilitation center for acquired brain injury survivors, where he helped start an evidence-based program for building psychological resilience during the COVID-19 pandemic.

Bhattacharyay is currently pursuing an MD at Harvard Medical School with aspirations of becoming a physician-engineer in neurocritical or neurosurgical care. At Harvard, he is researching sources of bias in medical AI to protect patient safety and equity in the clinical deployment of decision support systems for TBI care. Bhattacharyay's mission is to enhance the precision and global accessibility of TBI care with big data.

Min Jae Kim, Engr '22

Min Jae immigrated from Korea to Fairfax, Virginia, when he was 14. He completed his undergraduate education at Johns Hopkins University in biomedical engineering and neuroscience.

As a college student, Kim became interested in studying underlying brain circuit dynamics and how selectively intervening in this circuitry through neuromodulatory therapies can improve clinical outcomes in movement disorders and epilepsy. He worked closely with Kelly Mills , director of the Movement Disorders Division and an associate professor of neurology at Johns Hopkins, to identify neural circuitry associated with cognitive impairment in patients with Parkinson's disease after deep brain stimulation. Additionally, he collaborated with Johns Hopkins neurologist Joon-Yi Kang and neurosurgeon William Stanley Anderson to investigate radiographic markers and circuits to enhance seizure freedom rates for epilepsy patients undergoing minimally invasive epilepsy surgery. From this work, Kim has held a patent as a lead inventor and was named a Barry Goldwater Scholar in 2021 .

After completing his undergraduate degree, Kim pursued additional training in understanding neural circuitry in movement disorders and neuropsychiatric disorders with Andreas Horn at Network Stimulation Laboratory and Harvard Medical School before beginning his MD/PhD training at the University of Pennsylvania. Throughout his training, Kim's goal has been to study circuit-level pathophysiology in neurological disorders and translate his research findings to revolutionize the clinical landscape of neuromodulation. He is currently investigating novel methods to optimize neuromodulatory therapies across numerous neurological and neuropsychiatric disorders at Penn Medicine and Children's Hospital of Philadelphia alongside multidisciplinary research and clinical faculty members, including professors Casey Halpern, Kathryn Davis, Benjamin Kennedy, Han-Chiao Isaac Chen, and Iahn Cajigas.

Kim has published more than 18 papers in many reputable journals such as Biological Psychiatry , Epilepsia , Neurosurgery , and Brain Stimulation . His research works have been recognized by both national and international organizations such as the American Epilepsy Society, International Parkinson and Movement Disorder Society, and the Congress of Neurological Surgeons. As a future neurosurgeon-scientist, he aims to develop next-generation neuromodulatory therapeutics to repair neurophysiological and network dysfunctions in neurological disorders.

James Occean

James Occean emigrated from Haiti to the U.S. at the age of 10. He later pursued a bachelor of science degree in biomedical sciences at the University of South Florida as a first-generation college student. Under the mentorship of Abraham Salinas-Miranda and Nicholas Thomas, he conducted epidemiological research to identify predictors and risk factors for intimate partner violence among women in his native country, Haiti. This effort culminated in James' first lead-author publication in the Journal of Interpersonal Violence. James then expanded his research into the biological sciences to understand how trauma exposure increases susceptibility to psychiatric disorders, a phenomenon typically observed in trauma-exposed women in Haiti. He joined Monica Uddin's lab and studied genetic and epigenetic mechanisms that underlie PTSD. His first-author publication in Psychiatry Research revealed that DNA methylation at a stress-sensitive gene influences the likelihood of developing PTSD after experiencing certain traumas.

After completing his undergraduate studies, James received the post-baccalaureate IRTA fellowship from the National Institute on Aging, National Institutes of Health. In Payel Sen's lab, he investigated how changes in epigenetic modifications and chromatin drive mammalian aging and related decline. During his two years in the Sen lab, James led and contributed to several peer-reviewed publications, secured over $140,000 in research grants for his work on DNA hydroxymethylation, and received the Early Career Scholar award from the American Aging Association.

Following his fellowship, James on track to earn his master's in bioinformatics at Johns Hopkins University in May. Concurrently, he works as a data scientist at Personal Genome Diagnostics within Labcorp Oncology, where he actively contributes to the verification and validation of noninvasive diagnostic assays designed to detect cancer-related and clinically relevant genetic alterations.

This fall, he will begin his PhD in Cancer Biology at Stanford. There, he plans to explore the genetic and epigenetic mechanisms driving tumor initiation, progression, and treatment resistance. His goal is to use this research to develop noninvasive cancer technologies and identify potential therapeutic targets.

Michael E. Xie

Michael E. Xie was born in New Jersey to immigrants from China, who came to the United States to pursue educational opportunities. As a child, Xie spent time living with his extended family in Jiangxi and Zhejiang provinces. Xie graduated from Harvard University summa cum laude and Phi Beta Kappa with a bachelor's degree in chemistry and physics and concurrent master's degree in statistics. As an undergraduate, he conducted research with Adam Cohen and developed an interest in neuroscience. In the lab, Xie was captivated by the modern ability to record detailed electrical signals from many individual neurons simultaneously, and he collaborated with Liam Paninski's group at Columbia University to develop new statistical tools that enable accurate interpretation of such recordings. His research resulted in a first-author publication in Cell Reports as well as co-authored publications in Nature and Cell . His undergraduate thesis also won a Thomas Temple Hoopes Prize from Harvard.

Currently, Xie is pursuing an MD/PhD in the Medical Scientist Training Program at Johns Hopkins University School of Medicine and Department of Biomedical Engineering and anticipates earning his degree in 2028. His PhD research, co-advised by Karel Svoboda and Adam Charles , uses novel neural recording techniques to examine the fundamental—but unanswered—question of what computations the individual neurons that make up the living brain can perform. With these insights, Xie hopes to build improved computational models of the brain that can help us understand how cognitive function may deteriorate with neuropsychiatric or neurodegenerative disease. Xie also leads a neurosurgery research project in the lab of Risheng Xu , assistant director, of the neurosurgery residency program, building deep learning models to improve patient outcomes.

To learn more about applying for the Soros Fellowship and other scholarships, visit the university's National Fellowship Program website .

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Tagged alumni , fellowships , graduate students

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