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Alpha 1 Antitrypsin Role in Health and Disease 1st Edition Adam Wanner
University of Hertfordshire Hatfield, Hertfordshire, UK
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Alpha-1Antitrypsin
Methods and Protocols
Edited by Cynthia L. Bristow
Alpha-1 Biologics, Long Island High Technology Incubator, Stony Brook University, Stony Brook, NY, USA
Editor Cynthia L. Bristow
Alpha-1 Biologics, Long Island
High Technology Incubator
Stony Brook University
Stony Brook, NY, USA
ISSN 1064-3745ISSN 1940-6029 (electronic)
Methods in Molecular Biology
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Preface
During the previous 30 years, it has become clear that the function of Alpha-1 Antitrypsin (AAT, alpha-1 proteinase inhibitor, α1PI) extends beyond simply inhibition of proteinases to its more significant functions involving cell signaling and cellular locomotion. The influence of AAT on the immune system has long been recognized, yet the exact nature of the influence was largely unknown other than its increased concentration in circulation during the acute phase of inflammation and its inhibition of some proteinases including neutrophil elastase which is known to damage tissue when left unchecked by AAT.
Active AAT circulates in blood in two isoforms in dynamic equilibrium: (1) native AAT, which binds irreversibly to neutrophil elastase, and (2) thiol-modified AAT, which binds reversibly to neutrophil elastase. Inactive AAT arises during infection or inflammation via modification of active AAT by factors released from microorganisms or host cells. Inactive AAT can arise by covalently complexing with soluble or cell-surface neutrophil elastase, being cleaved by proteinases other than elastase, by oxygenation, and by antibodies that have specificity for HIV envelope protein gp120. In its inactivated form, AAT binds to low density lipoprotein (LDL), apoB100, and members of the LDL receptor family (LDL-RFMs) whereby AAT facilitates LDL uptake into cells.
Much of the progress in learning the unique AAT functions grew from advances in recognizing the various isoforms of AAT, measuring their specific activities, and discovering their effects on cellular activities. However, most investigations on the effects of AAT and experimental outcomes continue to quantify AAT in terms of protein concentration rather than relating those outcomes to AAT isoforms and specific activities thereby sometimes leading to misinterpretation and wrong turns. Because of increased knowledge of the crucial functions of AAT in innate immunity and the need to better understand its primordial importance throughout evolution, this collection of protocols is intended to elevate interest in further exploring the multiple, fascinating activities of AAT isoforms in innate immunity.
Stony Brook, NY, USACynthia L. Bristow
Alisha M. Gruntman, Wen Xue, and Terence R. Flotte
2 Approaches to Therapeutic Gene Editing in Alpha-1 Antitrypsin Deficiency
Alisha M. Gruntman, Wen Xue, and Terence R. Flotte
3 Manipulation of Proteostasis Networks in Transgenic ZAAT Zebrafish via
Connie Fung, Lee B. Miles, Robert J. Bryson-Richardson, and Phillip I. Bird 4
Kalsheker 5 Native and Ion Mobility Mass Spectrometry Characterization of Alpha 1 Antitrypsin Variants and Oligomers 41
Sarah Vickers, James Irving, David A. Lomas, and Konstantinos Thalassinos
6 Sanger and Next-Generation Sequencing of AAT
Valentina Barzon, Ilaria Ferrarotti, and Stefania Ottaviani
7 Measuring the Concentration of Active Alpha-1 Antitrypsin in Serum
Cynthia L. Bristow
8 Characterization of Novel Alpha-1-Antitrypsin Coding Variants in a Mammalian Cellular Model
Andrea Denardo, Emna Ben Khlifa, Mattia Bignotti, and Annamaria Fra 9 Alpha-1-Antitrypsin (A1AT) Proteotyping by LC-MS/MS
Jennifer Kemp, Paula M. Ladwig, and Melissa R. Snyder 10 Serum Western Blot for the Detection of a c-Myc Protein Tag in Non-human Primates and Mice
Meghan Blackwood, Qiushi Tang, and Alisha M. Gruntman
11 An Enzyme-Linked Immunosorbent Assay (ELISA) for Quantification of Circulating Pi*Z Alpha1-Antitrypsin Polymers .
Sabina Janciauskiene, Tobias Welte, and Matthias Lehmann
12 Measuring of Alpha-1 Antitrypsin Concentration by Nephelometry or Turbidimetry .
Carmen Marin-Hinojosa, Daniel Fatela-Cantillo, and Jose Luis Lopez-Campos
13
Autoantibody Recognition of Natural and Homocysteinylated Alpha-1 Antitrypsin: Indirect Enzyme-Linked Immunosorbent Assay (ELISA) Quantification in Sera
Tania Colasanti and Fabrizio Conti
14 Monitoring the Secretion and Activity of Alpha-1 Antitrypsin in Various Mammalian Cell Types 143
Kevin P. Guay, Haiping Ke, Lila M. Gierasch, Anne Gershenson, and Daniel N. Hebert
PART III AAT INTERACTIONS WITH LIPOPROTEINS AND LIPOPROTEIN RECEPTORS
15 Lipoproteins in Negative Feedback with Alpha-1 Antitrypsin 167
Cynthia L. Bristow
16 Silencing Very-Low-Density Lipoprotein Receptor Reveals Alpha-1 Antitrypsin Role in HIV Infectivity
Cynthia L. Bristow
175
17 Purified Versus Plasma Alpha-1 Antitrypsin Effects on Cellular Activities . . . . . . 185
Cynthia L. Bristow
Index
Contributors
VALENTINA BARZON • Department of Internal Medicine and Therapeutics, Pulmonology Unit, University of Pavia, Pavia, Italy
EMNA BEN KHLIFA • Department of Molecular and Translational Medicine, University of Brescia, Brescia, Italy
MATTIA BIGNOTTI • Department of Molecular and Translational Medicine, University of Brescia, Brescia, Italy
PHILLIP I. BIRD • Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, VIC, Australia
MEGHAN BLACKWOOD • Department of Pediatrics and Horae Gene Therapy Center, University of Massachusetts Medical School, Worcester, MA, USA
CYNTHIA L. BRISTOW • Alpha-1 Biologics, Long Island High Technology Incubator, Stony Brook University, Stony Brook, NY, USA
ROBERT J. BRYSON-RICHARDSON • School of Biological Sciences, Monash University, Clayton, VIC, Australia
TANIA COLASANTI • Rheumatology Unit, Department of Clinical Internal, Anesthesiological and Cardiovascular Sciences, Sapienza University of Rome, Rome, Italy
FABRIZIO CONTI • Rheumatology Unit, Department of Clinical Internal, Anesthesiological and Cardiovascular Sciences, Sapienza University of Rome, Rome, Italy
ANDREA DENARDO • Department of Molecular and Translational Medicine, University of Brescia, Brescia, Italy
DANIEL FATELA-CANTILLO • Unidad de Gestion Clı ´ nica de Laboratorios, Servicio de Bioquı ´ mica Clı ´ nica, Seccion de Inmunoproteı ´ nas y Marcadores Tumorales, Hospital Universitario Virgen del Rocı ´ o/Universidad de Sevilla, Seville, Spain
ILARIA FERRAROTTI • Centre for Diagnosis of Inherited Alpha-1 Antitrypsin Deficiency, UOC Pulmonology, Fondazione IRCCS Policlinico San Matteo, Pavia, Italy
TERENCE R. FLOTTE • Department of Pediatrics, University of Massachusetts Chan Medical School, Worcester, MA, USA
ANNAMARIA FRA • Department of Molecular and Translational Medicine, University of Brescia, Brescia, Italy
CONNIE FUNG • Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, VIC, Australia
ANNE GERSHENSON • Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA; Program in Molecular and Cellular Biology, University of Massachusetts, Amherst, MA, USA
LILA M. GIERASCH • Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA; Program in Molecular and Cellular Biology, University of Massachusetts, Amherst, MA, USA; Department of Chemistry, University of Massachusetts, Amherst, MA, USA
ALISHA M. GRUNTMAN • Department of Pediatrics, University of Massachusetts Chan Medical School, Worcester, MA, USA; Department of Pediatrics and Horae Gene Therapy Center, University of Massachusetts Medical School, Worcester, MA, USA
KEVIN P. GUAY • Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA; Program in Molecular and Cellular Biology, University of Massachusetts, Amherst, MA, USA
DANIEL N. HEBERT • Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA; Program in Molecular and Cellular Biology, University of Massachusetts, Amherst, MA, USA
JAMES IRVING • Institute of Structural and Molecular Biology, Birkbeck College, University of London, London, UK; UCL Respiratory, University College London, London, UK
SABINA JANCIAUSKIENE • Department of Respiratory Medicine, Hannover Medical School, Biomedical Research in Endstage and Obstructive Lung Disease Hannover (BREATH), Member of the German Center for Lung Research (DZL), Hannover, Germany
NOOR KALSHEKER • University of Nottingham, Nottingham, UK
HAIPING KE • Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA
JENNIFER KEMP • Division of Clinical Biochemistry and Immunology, Department of Laboratory Medicine and Pathology, Mayo Clinic, Rochester, MN, USA
PAULA M. LADWIG • Division of Clinical Biochemistry and Immunology, Department of Laboratory Medicine and Pathology, Mayo Clinic, Rochester, MN, USA
DAVID A. LOMAS • UCL Respiratory, University College London, London, UK
JOSE LUIS LOPEZ-CAMPOS • Unidad Me´dico-Quiru ´ rgica de Enfermedades Respiratorias, Instituto de Biomedicina de Sevilla (IBiS), Hospital Universitario Virgen del Rocı ´ o/CSIC/ Universidad de Sevilla, Seville, Spain; CIBER de Enfermedades Respiratorias (CIBERES), Instituto de Salud Carlos III, Madrid, Spain
CARMEN MARIN-HINOJOSA • Unidad Me´dico-Quiru ´ rgica de Enfermedades Respiratorias, Instituto de Biomedicina de Sevilla (IBiS), Hospital Universitario Virgen del Rocı ´ o/CSIC/ Universidad de Sevilla, Seville, Spain; CIBER de Enfermedades Respiratorias (CIBERES), Instituto de Salud Carlos III, Madrid, Spain
LEE B. MILES • School of Biological Sciences, Monash University, Clayton, VIC, Australia
STEFANIA OTTAVIANI • Centre for Diagnosis of Inherited Alpha-1 Antitrypsin Deficiency, UOC Pulmonology, Fondazione IRCCS Policlinico San Matteo, Pavia, Italy
MELISSA R. SNYDER • Division of Clinical Biochemistry and Immunology, Department of Laboratory Medicine and Pathology, Mayo Clinic, Rochester, MN, USA
QIUSHI TANG • Department of Pediatrics and Horae Gene Therapy Center, University of Massachusetts Medical School, Worcester, MA, USA
KONSTANTINOS THALASSINOS • Institute of Structural and Molecular Biology, Division of Biosciences, University College London, London, UK; Institute of Structural and Molecular Biology, Birkbeck College, University of London, London, UK
SARAH VICKERS • Institute of Structural and Molecular Biology, Division of Biosciences, University College London, London, UK; Institute of Structural and Molecular Biology, Birkbeck College, University of London, London, UK
TOBIAS WELTE • Department of Respiratory Medicine, Hannover Medical School, Biomedical Research in Endstage and Obstructive Lung Disease Hannover (BREATH), Member of the German Center for Lung Research (DZL), Hannover, Germany
WEN XUE • Department of Pediatrics, University of Massachusetts Chan Medical School, Worcester, MA, USA
Alpha-1 Antitrypsin Deficiency
Alisha M. Gruntman, Wen Xue, and Terence R. Flotte
Abstract
Alpha-1 antitrypsin (AAT) deficiency is a common monogenic disorder in which there is a strong founder effect of a single missense mutation in SERPINA1, the gene encoding this major circulating serum antiprotease that is normally expressed primarily in hepatocytes. These features make AAT deficiency particularly attractive as a target for therapeutic gene editing using a wide variety of approaches.
1 Alpha-1 Antitrypsin Deficiency, the SERPINA1 Locus and Its Disease-Causing Alleles
1.1 Brief History
The familial clustering of emphysema cases led the pioneering clinical investigators Laurell and Eriksson to discover that the absence of the alpha-1 globulin band on total serum protein electrophoresis was directly correlated with the presence of lung disease in affected families [1, 2]. They further showed that the alpha-1 band possessed most of the serum biochemical activity to inactivate trypsin. The protein comprising this band was named alpha-1 antitrypsin (AAT) and the deficiency disease was called AAT deficiency (AATD) (Fig. 1). The subsequent discovery of the gene and its mutations set the stage for AATD to become one of the best studied single gene diseases [3].
2 Characterization of the Protein and Gene
Molecular studies of the AAT protein and the gene that encodes it have revealed that AAT is a 52 kD glycoprotein with 394 amino acids and 3 asparaginyl linked carbohydrate chains. It is a member of the serine proteinase inhibitor (SERPIN) gene family, along with
structurally similar proteins such as anti-thrombin 3 (AT3) [4]. Members of this family primarily function as proteinase inhibitors, and AAT has strong antiprotease activity against neutrophil elastase (NE), which is likely its most relevant target, along with anti-trypsin activity and activity against proteinase-3, certain cathepsins, metalloproteinases, and caspase-3. Physiologically, this enables AAT to be broadly anti-inflammatory and to function as a buffer against proteases and other pro-inflammatory molecules released within loci of infection. AAT is a secreted protein, with primary sites of synthesis being hepatocytes, monocytes, and macrophages.
Fig. 1 Absence of alpha-1 globulin with antitryptic activity band on serum protein electrophoresis. (Reproduced from Erikson, 1964 with permission from John Wiley and Sons, Inc)
AAT is a very abundant serum protein, with circulating plasma levels in humans typically ranging from 20 to 50 μM, making it the second most abundant serum protein after albumin. AAT levels increase during periods of fever, infection, and acute physiologic stress. Subsequently, it has been discovered that its synthesis is specifically induced by interleukin-6 (IL-6) during such episodes.
The structure of both wild-type and common mutant AAT proteins has been solved by X-ray crystallography. The protein structure is similar to other serpins with nine alpha helices, three beta pleated sheets, two internal salt bridges, and a less structured reactive loop [5, 6]. The reactive loop contains the sites that interact covalently with NE when AAT complexes with NE to inactivate it. Methionine residues in the active loop may be inactivated by oxidation. It is thought that the sensitivity of AAT to oxidative inactivation allows for NE to remain fully active at sites of intense inflammation, such as within bacterial abscesses, in which the release of reactive oxygen species from the neutrophil oxidative burst would be predicted to limit AAT function.
The most common AAT mutation, E342K, causes a change of charge valence of +2, a difference which makes this mutant protein easy to distinguish on isoelectric focusing gel (IEF) electrophoresis. The IEF appearance of wild-type and E342K AAT has led to their naming with letters as the various IEF bands were labeled. The wild-type was labeled “M” or proteinase inhibitor M (PiM), while the common E342K mutation was labeled as PiZ. Various other mutant proteins have been described based initially on IEF gel mobility and later from gene sequencing [7, 8].
3 The SERPINA1 Locus in Humans and Mice
The human SERPINA1 gene is spread over 12.2 kb of chromosome 14 q31–31.2. It consists of seven exons (IA,IB,IC, and II through V), three of which are alternative first exons, depending on the transcription start site. Thus, the most abundant mRNA species possesses only five exons. The start codon is in exon II followed by the ATG followed by a 24-amino-acid leader sequence.
The corresponding murine gene is found on murine chromosome 12. Complicating the murine genomic sequence (and the engineering of transgenic mice) is the fact that the gene is repeated in a tandem of isoforms. There are seven distinct isoform sequences, with the genomes of various strains of mice hosting between three and five distinct copies of Serpina1 on each allele [9].
4 Mutations and Founder Effects
At least 100 different AAT alleles and polymorphisms have been described [10]. The most frequent wild-type allele, M1(Val213), has an allele frequency of 0.44–0.49, while the next most common non-disease-causing alleles, M1(Ala213), M2, and M3, comprise most of the remaining 50% in the normal population. There is a strong founder effect for the most common mutant allele (PI*Z is the designation for the DNA mutation, PiZ is the mutant protein designation) in several parts of Europe, most notably in Scandinavia, Ireland, Spain, and other parts of Western Europe. The PI*Z mutation causes a change from a glutamate residue to a lysine residue in the protein (E342K). The PI*Z mutation most commonly occurs on an allele that also possesses the M1(Val213) sequence. The PiS missense mutation (G264V) is another common mutation that appears to be partially functional. PiSZ compound heterozygotes generally have intermediate serum levels and an intermediate phenotype. Studies of the natural history of such compound heterozygotes led to the conclusion that a plasma level of 11 μM AAT would be protective against lung disease [3, 11, 12].
5 Sequence Context for the Common PI*Z Mutation
The specifics of the PiZ mutation are important to consider in the context of gene editing, base editing, and prime editing. The linear sequence around the mutation site is shown in Fig. 2. Features relevant to the gene editing include a number of potential protospacer adjacent motif (PAM) sequences in the vicinity. For example, the original Streptococcus pyogenes-Cas9 (SpCas9) requires a PAM sequence of NGG, of which there are two instances near the mutation site, while other SpCas9-based mutants may have more liberal PAM requirements. The specifics of this relative to design of the necessary single guide RNAs (sgRNAs) for inducing double-strand breaks (DSBs) or base edits or for the design of prime-editing guide RNAs (pegRNAs) will be addressed below, but generally an sgRNA must include both PAM sequence specificity and base complementarity to the sequence to be targeted for endonuclease activity or other enzyme-mediated nucleotide alterations.
6 Consequence of Mutation and Disease Manifestations
AATD has been variously described as autosomal recessive or as co-dominant with variable penetrance. Patients with homozygous, severe deficiency of AAT are at high risk for spontaneously
Fig. 2 Wild-type (PI*M) and common mutation (PI*Z) DNA sequence SERPINA1 gene sequence at the site of the mutation
developing lung disease in early adulthood, while heterozygotes have a statistically increased risk of lung disease when exposed to tobacco smoking or other environmental insults.
The most common serious manifestation of AATD is lung disease, which can have a number of features. Clinically, patients often initially experience an asthma-like picture with wheezing and partially reversible airways obstruction. The chronic obstructive pulmonary disease (COPD) picture can progress with features including bronchiectasis, emphysema, and exacerbations of respiratory distress often triggered by intercurrent respiratory infections. The gradual nature of AATD progression and its similarity to non-Mendelian diseases like asthma, COPD, and idiopathic bronchiectasis often leads to delayed diagnosis, with most AATD lung disease patients being diagnosed in their 40s or 50s, in spite of years of preceding symptoms. Progression of lung disease may lead to the more classically described radiographic and histopathologic features of bibasilar emphysematous changes on chest X-ray and computed tomography (CT) scans and of panacinar emphysema on biopsy (Fig. 3)
Liver disease is a more puzzling manifestation of AATD. There is remarkable genetic homogeneity among the disease mutations, with PiZ comprising over 90% of disease mutations. Even among PiZ homozygotes, only 5% develop life-threating liver disease, but 30–50% of patients develop at least subclinical manifestations of liver injury at some time in infancy, childhood, or adulthood. The most severe forms of liver disease begin with significant inflammation and may progress to cholestasis, fibrosis, cirrhosis, and liver failure and may lead to hepatocellular carcinoma. The co-factors leading to such a variable liver presentation of PiZ homozygous patients remain unknown, but it does seem clear that AATD liver disease is due to a toxic gain of function of the PiZ mutant protein [4]. Thus, inactivation of the PiZ gene by editing could result in clinical benefit for the liver disease, even in the absence of restoration of PiM synthesis and secretion.
There are other manifestations of AATD that are even more rare. The progression of vasculitis in some patients can manifest in panniculitis, in which painful subcutaneous nodules form at foci at subcutaneous vasculitis. Such patients often have positive anti-neutrophil cytoplasmic antibodies (ANCA). In addition to the vasculitis risk, an increased risk of hepatocellular carcinoma has been mentioned.
The cr ystal structure of PiZ mutant AAT has been well characterized. The change in charge from -1 to +2 in a region of beta sheet disrupts an important salt bridge, making possible the insertion of the reactive loop from a neighboring AAT molecule between the beta sheet strands, in a process known as “loopsheet” polymerization. Both the nascent decrease in stability of the normally folded PiZ protein and the propensity of the mutant to polymerize impair its secretion. This leads to both accumulation of the mutant protein within hepatocytes and to severe deficiency of circulating AAT activity. It is thought that the lung disease seen in AATD patients derives from this serum deficiency, while the liver disease which occurs in a subset of patients is a result of the abnormal mutant protein within the hepatocytes. These molecular properties are important in devising distinct gene editing strategies for therapy for the lung disease as opposed to for the liver disease.
Fig. 3 Panacinar emphysema in AATD lung disease
References
1. Laurell CB, Eriksson S (2013) The electrophoretic α1-globulin pattern of serum in α1-antitrypsin deficiency. COPD 10(Suppl 1):3–8
2. Eriksson S (1964) Pulmonary emphysema and alpha1-antitr ypsin deficiency. Acta Med Scand 175:197–205
3. Crystal RG (1989) The alpha 1-antitrypsin gene and its deficiency states. Trends Genet 5: 411–417
4. Strnad P, Mcelvaney NG, Lomas DA (2020) Alpha(1)-antitr ypsin deficiency. N Engl J Med 382:1443–1455
5. Sosulski ML, Stiles KM, Frenk EZ et al (2020) Gene therapy for alpha 1-antitrypsin deficiency with an oxidant-resistant human alpha 1-antitrypsin. JCI Insight 5:e135951
6. Lomas DA, Evans DL, Finch JT et al (1992) The mechanism of Z alpha 1-antitrypsin accumulation in the liver. Nature 357:605–607
Renoux C, Odou MF, Tosato G et al (2018) Description of 22 new alpha-1 antitrypsin genetic variants. Orphanet J Rare Dis 13:161
8. Crowther DC, Belorgey D, Miranda E et al (2004) Practical genetics: alpha-1-antitrypsin deficiency and the serpinopathies. Eur J Hum Genet 12:167–172
9. Barbour KW, Wei F, Brannan C et al (2002) The murine alpha(1)-proteinase inhibitor gene family: polymorphism, chromosomal location, and structure. Genomics 80:515–522
10. Seixas S, Marques PI (2021) Known mutations at the cause of alpha-1 antitrypsin deficiency an updated overview of SERPINA1 variation spectrum. Appl Clin Genet 14:173–194
11. Wewers MD, Casolaro MA, Sellers SE et al (1987) Replacement therapy for alpha 1-antitrypsin deficiency associated with emphysema. N Engl J Med 316:1055–1062
12. De Serres FJ (2003) Alpha-1 antitrypsin deficiency is not a rare disease but a disease that is rarely diagnosed. Environ Health Perspect 111:1851–1854
Part I
Alpha-1 Antitrypsin Polymorphisms and Gene Exon Usage
Approaches to Therapeutic Gene Editing in Alpha-1 Antitrypsin
Deficiency
Alisha M. Gruntman, Wen Xue, and Terence R. Flotte
Abstract
Five distinct gene therapy approaches have been developed for treating AATD. These approaches include knockout of the mutant (PiZ) allele by introduction of double-strand breaks (DSBs) and subsequent creation of insertions and deletions (indels) by DSB repair, homology-directed repair (HDR) targeted to the mutation site, base editing, prime editing, and alternatively targeted knock-in techniques. Each approach will be discussed and a brief summary of a standard CRISPR–Cas9 targeting method will be presented.
Key words Double-strand breaks, Indels, Homology-directed repair, Base editing, Prime editing
1 Molecular Strategies for Editing the SERPINA1 Gene
Several distinct molecular approaches to AAT gene editing have been developed and evaluated at proof of concept. These generally rely on one of five different strategies, each involving a Cas9 nuclease or mutant Cas9 (dCas9) fused with other relevant enzyme domains required for base editing or prime editing (Table 1).
The five strategies include the following: (1) the induction of inser tions and deletions (indels) by way of introducing a DNA double-strand break (DSB) within the SERPINA1 gene and then relying on DSB repair via non-homologous end-joining (NHEJ), (2) homology-directed repair (HDR) of the SERPINA1 gene in which a DSB is introduced near the PI*Z mutation site within the SERPINA1 gene coupled with the transfection of a template DNA strand with a corrected sequence at the mutation site and homology arms on either side, (3) a similar HDR approach instead of targeting a sequence just downstream of a very powerful promoter and using an HDR template with a complete wild-type PI*M cDNA coding sequence flanked by homology arms spanning the desired insertion site (so-called promoter hijacking), (4) a base
Introduction of indels to inactive endogenous PI*Z mutant allele
Induction of HDR with CRISPR–Cas9 to correct PI*Z locus
Use of HDR induced by CRISPR–Cas9 to knock-in a PI*M coding sequence downstream of a strong promoter (promoter hijacking)
Base editing to reverse mutation without inducing DSBs
Prime editing to rewrite mutation site with reverse transcriptase
editing approach in which a mutant Cas9 (dCas9) is fused at the protein level with a specialized adenosine deaminase (ADAR) domain to create an adenine base editor (ABE) capable of reversing the G-to-A nucleotide change on the sense strand at the PI*Z mutation site or a cytosine base editor (CBE) to alter the complementary strand, and (5) a prime editing approach utilizing a different dCas9 fusion, this one to reverse transcriptase (RT), which can then rewrite any sequence of appropriate length encoded by a template at the 3′ end of the pegRNA (Fig. 1).
1.1 Knockout
Expression of Mutant PiZ AAT Protein by NHEJ to Introduce Insertions and Deletions (Indels) in the Endogenous PI*Z Sequence
In a circumstance in which the only goal was to prevent or reverse AATD liver disease, the ability to knock out the expression of the endogenous mutant PI*Z gene would be desirable. Using CRISPR–Cas9 to introduce DSB into the DNA of the PI*Z locus would then be sufficient to induce NHEJ-mediated repair, a process which most commonly creates indels of 1–2 nucleotides, thereby causing a frameshift mutation which usually inactivates gene expression [1, 2]. For this purpose, the design of the sgRNA is critical. As noted above, the sgRNA must include a PAM sequence compatible with the Cas9 enzyme in use at the time. A table of the PAM specificities of the various native Cas9 enzymes available is provided below (Table 2).
Please note that mutant versions of the commonly employed Cas9s have been developed to change the PAM specificity. It is often helpful to utilize a bioinformatic sgRNA design program, several of which are publicly available [3]. As depicted in Fig. 1, once an appropriate sgRNA design is identified, active Cas9 endonuclease will create a double-strand break (DSB) at the site, which will then enable one of two mechanisms for site-specific gene editing.
Fig. 1 CRISPR-based mechanisms of homology-directed repair (HDR) and non-homologous end-joining (NHEJ) after introduction of DNA double-strand breaks (DSBs). (This figure was created using BioRender)
Table 2
PAM specificity of commonly used Cas9 enzymes
2 Methods
Design and validate CRISPR reagents targeting the AAT sequence [4]:
1. Input AAT genomic sequence using tools, such as https:// portals.broadinstitute.org/gppx/crispick/public.Alternatively, sgRNA can be manually selected by choosing 20-bp sequence upstream of NGG PAM.
2. Order oligonucleotides for sgRNA cloning.
3. Clone sgRNA into an sgRNA expression construct (e.g., pX330 Addgene #42230). The construct co-expresses Cas9 and sgRNA.
4. Transfect plasmids into 293 T cells.
5. After 3 days, collect genomic DNA and PCR amplify AAT region.
6. Perform T7E1, Sanger sequencing, or targeted deep sequencing to quantify indel rate.
3 HDR-Mediated Correction
The more common approach to the utilization of DSB creation with CRISPR–Cas9 is to utilize this DSB introduction to enhance homology-directed repair (HDR). With this approach, one utilizes a single-stranded or double-stranded DNA donor with homology arms flanking a corrected DNA sequence, along with the Cas9 and the sgRNA. The Cas9-endonuclease-mediated DSB-based mechanisms are demonstrated in the figure below.
We and others showed that Cas9-mediated HDR can partially correct the human PiZ mutation in a widely used transgenic mouse model of AATD (Fig. 1)[5]. The PiZ transgenic mice express high levels of human Z-AAT, most of which is sequestered in hepatocytes [6]. We reported that adeno-associated virus (AAV) delivery of Cas9, sgRNA, and an HDR template corrected the PiZ mutation in the adult mouse liver and restored M-AAT to ~8% of the level required to improve lung function [5].
The small size of saCas9 allows sgRNA and Cas9 to be packaged in one AAV vector. Shen et al. showed that AAV8-saCas9 targeting exon 2 of PiZ leads to reduced PiZ aggregates in mouse hepatocytes [7]. When sgRNA was designed to target exon 5 and a second AAV was used to deliver HDR donor, the treated mice had ~5% M-AAT mRNA restored. Together, this work is proof of principle that CRISPR can correct the PiZ mutation in vivo.
Despite this success, HDR requires co-delivery of a long DNA donor and works poorly in non-dividing or slow-dividing cells. These limitations of HDR can be addressed using base editing or prime editing.
4 Knock-in of cDNA
The knock-in approach utilizes HDR to introduce a full-length wild-type PI*M SERPINA1 coding sequence downstream of an active promoter that is highly expressed within hepatocytes. There are two versions of this approach that have been published for AATD. This approach has been used in a recent publication in which a recombinant adenovirus vector was used to introduce the
Cas9 and sgRNA and an HDR template with homology arms matching the ROSA26 locus, flanking a full expression cassette with an active promoter, full SERPINA1 cDNA, and polyadenylation signal [8]. The ROSA26 locus has been used for several similar targeted integration approaches because it represents a so-called genomic safe harbor (GSH) where integrations generally are expressed but do not seem to induce the expression of latent proto-oncogenes. The AAVS1 locus is another GSH commonly used for such purposes.
Another version of a full cDNA knock-in involves a so-called “promoter-less” or “promoter hijacking” approach. This approach was first applied by Barzel et al. to the delivery of a factor IX cDNA to the albumin locus, inserting the cDNA just downstream of the albumin promoter [9]. The introduction of a 2A element allows for normal expression of endogenous albumin from the same transcript. Interestingly, this was adapted for use in the context of an rAAV construct without exogenous Cas nuclease. Borel et al. utilized a similar construct targeted to the albumin locus, expressing PiM-AAT protein from the albumin promoter [10]. The Borel construct combined this augmentation approach with an allelespecific knockdown of the endogenous PiZ-AAT. These studies demonstrated that the expression from the albumin promoter was highly efficient within the subset of transduced hepatocytes and that these hepatocytes demonstrated a selective advantage for proliferation, regeneration, and repopulation of the liver.
5 Base Editing
Base editing utilizes a base editor enzyme comprising a Cas9 nickase fused to deaminase [11–13]. Cytosine base editor (CBE) is a fusion of cytidine deaminase and uracil glycosylase inhibitor to Cas9D10A nickase. Adenine base editor (ABE) comprises an evolved TadA adenosine deaminase fused to Cas9 nickase. CBEs and ABEs convert C to T or A to G, respectively.
Packer et al. applied CBE to install a compensatory Met374Ile mutation for PiZ and ABE to directly correct the PiZ mutation [14]. PiZ-transgenic mice were treated with lipid nanoparticles formulated with base editor mRNA and sgRNA. The treated mice exhibited up to 30% of correction of PiZ in the liver and increased serum AAT. CBE or ABE could mutate another C or A near the target region (bystander editing). This study reported that corrected AAT with common bystander mutations still inhibit elastase activity [14]. Recently, a report from the laboratory of Andrew Wilson has indicated that the potential for base editing may be more efficiently fulfilled in induced pluripotent stem cell (iPSC)derived hepatocytes [15]. This opens the possibility of ex vivo base editing followed by hepatocyte engraftment.
6 Prime Editing
PE is composed of Cas9 nickase fused to an engineered reverse transcriptase (RT) (Fig. 2)[16]. A prime editing guide RNA (pegRNA) targets the PE to the genomic target (Fig. 1). The pegRNA harbors a primer binding site (PBS) and RT template (RTT) for RT to copy the new genetic sequence from RTT into the target genomic site (Fig. 2)[16]. Compared with CRISPRHDR, PE does not induce double-strand breaks (DSBs) or HDR donor DNA. PE is also more flexible and produces fewer off-target editing than base editing [17]. PE (~6.3 kb) exceeds the packaging limit of AAV and requires dual AAV for delivery. We have recently reported an optimized PE (PE2*) that can correct PiZ mutations in the mouse liver [18]. Dual adeno-associated virus (AAVs) of a splitintein PE2* enables the correction of ~3% PiZ alleles in mouse liver. This work demonstrates the broad potential of this genome editing technology for disease gene correction in vivo.
References
1. Ijaz F, Nakazato R, Setou M et al (2022) A pair of primers facing at the double-strand break site enables to detect NHEJ-mediated indel mutations at a 1-bp resolution. Sci Rep 12: 11681
2. Guo T, Feng YL, Xiao JJ et al (2018) Harnessing accurate non-homologous end joining for efficient precise deletion in CRISPR/Cas9mediated genome editing. Genome Biol 19: 170
3. Brazelton VA Jr, Zarecor S, Wright DA et al (2015) A quick guide to CRISPR sgRNA design tools. GM Crops Food 6:266–276
4. Ran FA, Hsu PD, Wright J et al (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8:2281–2308
5. Song CQ, Wang D, Jiang Tet al (2018) In vivo genome editing partially restores alpha1antitrypsin in a murine model of AAT deficiency. Hum Gene Ther 29:853–860
6. Carlson JA, Rogers BB, Sifers RN et al (1989) Accumulation of PiZ alpha 1-antitrypsin causes liver damage in transgenic mice. J Clin Invest 83:1183–1190
7. Shen S, Sanchez ME, Blomenkamp K et al (2018) Amelioration of alpha-1 antitrypsin deficiency diseases with genome editing in transgenic mice. Hum Gene Ther 29:861–873
8. Stephens CJ, Kashentseva E, Everett W et al (2018) Targeted in vivo knock-in of human alpha-1-antitrypsin cDNA using adenoviral
Fig. 2 Design of Prime Editor and pegRNA. (This figure was created using BioRender)
delivery of CRISPR/Cas9. Gene Ther 25:139–156
9. Barzel A, Paulk NK, Shi Y et al (2015) Promoterless gene targeting without nucleases ameliorates haemophilia B in mice. Nature 517: 360–364
10. Borel F, Tang Q, Gernoux G et al (2017) Sur vival advantage of both human hepatocyte xenografts and genome-edited hepatocytes for treatment of α-1 antitrypsin deficiency. Mol Ther 25:2477–2489
11. Gaudelli NM, Komor AC, Rees HA et al (2017) Programmable base editing of A Tto G C in genomic DNA without DNA cleavage. Nature 551:464–471
12. Komor AC, Kim YB, Packer MS et al (2016) Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature 533:420–424
13. Hess GT, Tycko J, Yao D et al (2017) Methods and applications of CRISPR-mediated base editing in eukaryotic genomes. Mol Cell 68: 26–43
14. Packer MS, Chowdhary V, Lung G et al (2022) Evaluation of cytosine base editing and adenine base editing as a potential treatment for alpha-1 antitr ypsin deficiency. Mol Ther 30:1396–1406
15. Werder RB, Kaserman JE, Packer MS et al (2021) Adenine base editing reduces misfolded protein accumulation and toxicity in alpha-1 antitrypsin deficient patient iPSC-hepatocytes. Mol Ther 29:3219–3229
16. Anzalone AV, Randolph PB, Davis JR et al (2019) Search-and-replace genome editing without double-strand breaks or donor DNA. Nature 576:149–157
17. Habib O, Habib G, Hwang GH et al (2022) Comprehensive analysis of prime editing outcomes in human embryonic stem cells. Nucleic Acids Res 50:1187–1197
18. Liu P, Liang SQ, Zheng C et al (2021) Improved prime editors enable pathogenic allele correction and cancer modelling in adult mice. Nat Commun 12:2121
Manipulation of Proteostasis Networks in Transgenic ZAAT
Zebrafish via CRISPR–Cas9 Gene
Editing
Connie Fung, Lee B. Miles, Robert J. Bryson-Richardson, and Phillip I. Bird
Abstract
The CRISPR–Cas9 genome editing system is used to induce mutations in genes of interest resulting in the loss of functional protein. A transgenic zebrafish α1-antitrypsin deficiency (AATD) model displays an unusual phenotype, in that it lacks the hepatic accumulation of the misfolding Z α1-antitrypsin (ZAAT) evident in human and mouse models. Here we describe the application of the CRISPR–Cas9 system to generate mutant zebrafish with defects in key proteostasis networks likely to be involved in the hepatic processing of ZAAT in this model. We describe the targeting of the atf6a and man1b1 genes as examples.
Key words CRISPR–Cas9, Zebrafish, α1-antitrypsin, AATD, Atf6a, Man1b1
1 Introduction
The Z allele of α1-antitrypsin (ZAAT) predisposes individuals homozygous for this allele (Pi*ZZ) to lung damage due to insufficient levels of circulating α1-antitrypsin, and to liver disease due to misfolding and retention of the protein in hepatocytes [1]. Investigation of protein quality control factors such as ER chaperones and E3 ligases has demonstrated that the disruption of proteostasis networks in the liver can increase intracellular retention of ZAAT, leading to elevated hepatic stress and disease progression [2–7].
The transgenic zebrafish model expressing human ZAAT is a unique tool for ZAAT studies because hepatic inclusion bodies do not form despite animals displaying serum α1-antitrypsin insufficiency [8]. Understanding the molecular mechanism behind the efficient disposal of misfolded ZAAT by zebrafish hepatocytes may be useful in developing strategies to prevent the development of liver disease in Pi*ZZ patients.
Zebrafish are an attractive model system to study α1-antitrypsin deficiency (AATD) because animals are genetically outbred, so any confounding influence of unknown genetic mutations/modifiers is minimized. Indeed, the zebrafish model may better reflect the diverse genetic background of human AATD patients compared to inbred mouse models. The full zebrafish genome has been sequenced and annotated and exhibits high genomic similarity to humans. Over 80% of human disease-related genes have orthologs in the zebrafish genome [9].
The zebrafish embryo possesses a fully formed liver by 5 days post-fertilization and is transparent, allowing easy genetic manipulation and visualization of labeled tissues via fluorescence imaging. The low cost of maintenance and high fecundity of zebrafish allows the establishment of transgenic and mutant lines in a short timeframe, thus providing power for large-scale genetic studies. Furthermore, the easy delivery of chemicals to fish via surrounding water offers the potential of pursuing high-throughput drug screens.
One challenge of working with the zebrafish model is that many human genes have multiple corresponding homologs in the zebrafish genome due to a whole genome duplication event that occurred in the teleost linage after the divergence from the last common ancestor shared with humans, approximately 440 million years ago [10–12]. In order to accurately assess the influence protein quality control factors have on ZAAT processing, the zebrafish ortholog of the human gene being modeled must be identified. If this is unclear, because multiple homologs exist in the zebrafish genome, all corresponding zebrafish homologs to the factor of interest must be considered as part of the investigation (see Note 1). As described below, homologs can be easily and simultaneously targeted via gene editing.
The CRISPR–Cas9 genome editing system is widely used for gene manipulation in model organisms. The Cas9 endonuclease, when forming a ribonucleoprotein (RNP) complex with a short guide RNA (gRNA) sequence, can target DNA with high specificity to induce a double-stranded break [13]. Errors during the subsequent DNA repair process can cause small insertions or deletions (indel) in the genes targeted, resulting in frameshifts, premature stop codons, and loss of functional protein.
Here we describe the procedures to rapidly generate mutant zebrafish using CRISPR–Cas9-mediated gene editing. Briefly, zebrafish embryo is injected with the RNP complex, raised to adulthood, and out-crossed to wildtype fish. Mutations in heterozygous offspring are identified by PCR and Sanger sequencing. In the examples below, atf6a, which is the (single) ortholog of human ATF6, is targeted by injecting a single gRNA. We also describe the use of multiple CRISPR gRNAs to simultaneously target the man1b1a and man1b1b genes, which are homologs of human
2 Materials
2.1 Zebrafish Embryo Microinjection
MAN1B1. In our hands, using the components of the Alt-R® CRISPR–Cas9 system from Integrated DNA Technologies (IDT) in the modified protocol described below, gene editing is observed in 100% of zebrafish embryos following successful RNP delivery (see Subheading 3.5).
2.2 DNA Extraction + PCR
1. Adult ZAAT transgenic zebrafish: Tg(lfabp: eGFP-T2AZAAT) (see Note 2).
2. Nuclease-free IDTE buffer (IDT): 10 mM Tris-HCl, 0.1 mM EDTA. Store at -20 °C.
3. Nuclease-free duplex buffer (IDT): 30 mM HEPES, pH 7.5; 100 mM potassium acetate. Store at -20 °C.
4. 100 μM Alt-R® CRISPR–Cas9 crRNA (IDT): resuspend 2 nmol lyophilized powder in 20 μL nuclease-free IDTE buffer. Store at -20 °C(see Note 3).
5. 100 μM Alt-R® CRISPR–Cas9 TracrRNA (IDT): resuspend 5 nmol lyophilized powder in 50 μL nuclease-free IDTE buffer. Store at -20 °C.
6. Alt-R® S.p. HiFi Cas9 Nuclease V3 (61 μM) (IDT). Store at20 °C.
7. 1 M KCl: sterilize solution by passing through a 0.2 μm filter.
8. 0.5% (w/v) phenol red solution: sterilize solution by passing through a 0.2 μm filter.
9. 0.5% (w/v) cascade blue solution: store at -20 °C.
10. E3 embryo medium: 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM, and MgSO4 in distilled water.
11. Agarose powder.
12. Zebrafish injection mold (see Note 4).
13. Microinjector.
14. Microcapillary tips (Eppendorf Microloader).
15. 1 mm glass capillary, borosilicate thin wall with filament, 1.0 mm outer diameter, 0.78 mm inner diameter, 100 mm in length.
16. Micropipette puller.
17. Heat block at 37 °C.
1. 50 mM NaOH solution.
2. 1 M Tris-HCl pH 7.4.
3. Primers flanking the CRISPR target site(s) (see Note 5).
4. PCR master mix (see Note 6).
2.3 TBE-Acrylamide Gel
5. Nucleic acid dye (see Note 7).
6. Tricaine: 0.04% (w/v) ethyl 3-aminobenzoate methanesulfonate salt in Tris-HCl, pH 7.5. Store at 4 °C. Further dilute 1:10 (final concentration of 0.004%) into E3 embryo medium to facilitate anesthesia.
7. Scalpel blade.
8. Tweezers.
9. Vortex.
10. Heating block at 95 °C.
1. 30% acrylamide, 0.8% bis-acrylamide solution.
2. 2× TBE buffer: dissolve 43.2 g Tris-HCl, 22 g boric acid, and 2.32 g EDTA in 4 L distilled water.
3. 10% (w/v) ammonium persulfate.
4. Tetramethylethylenediamine (TEMED).
3 Methods
3.1 Preparing the Day Before Embryo Injection
3.2 Preparing Injection Mix
1. Place up to 15 fish into a large breeding tank with a divider separating males and females in the tank and leave overnight.
2. To prepare the injection plate, dissolve 1 g of agarose into 100 mL distilled water by carefully warming in a microwave. Pour the solution into a 10 cm petri dish until the solution reaches ~0.5 cm in height. Keep the remaining molten agarose at 60 °C in a water bath. Allow the agarose in the plate to set completely, then pour on more molten agarose until it reaches ~0.3 cm further in height. Immediately place the injection plastic mold (Fig. 1a) into the agarose in the petri dish, avoiding introducing bubbles in the troughs. Allow the agarose to set completely before removing the mold. The injection plate is now ready for use, or it can be stored at 4 °C for several months.
1. To prepare 30 μM gRNA complex solution (see Note 8), add 3 μL 100 μM CrRNA, 3 μL 100 μM TracrRNA, and 4 μL nuclease-free duplex buffer in a 0.2 mL microfuge tube, mix well, and incubate at 95 °C for 5 min.
2. To prepare 5 μL injection mix, add 1 μL gRNA complex (see Note 9), 0.35 μL Cas9 protein (700 pg/nL) (see Note 10), 1.5 μL 1 M KCl (300 mM) (see Note 11), 0.5 μL phenol red solution, 0.5 μL cascade blue solution, and 1.15 μL ultrapure water in a 0.2 mL microfuge tube, mix well, and incubate at 37 °C for 10 min to allow RNP complex formation. Allow the mixture cool to room temperature before injection.
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