engineering affinity and specificity of antibodies
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University of ConnecticutOpenCommons@UConn
Doctoral Dissertations University of Connecticut Graduate School
6-28-2019
Engineering Affinity and Specificity of AntibodiesTargeting Protein ModificationsDan LiUniversity of Connecticut - Storrs, [email protected]
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Recommended CitationLi, Dan, "Engineering Affinity and Specificity of Antibodies Targeting Protein Modifications" (2019). Doctoral Dissertations. 2226.https://opencommons.uconn.edu/dissertations/2226
Engineering Affinity and Specificity of Antibodies Targeting Protein Modifications
Dan Li, PhD
University of Connecticut, 2019
Antibodies targeting protein post-translational modifications (PTMs) are an essential tool in
scientific research and clinical investigations. Due to the transient and heterogeneous nature of
protein PTMs, high specificity and affinity of PTM-specific antibodies are critical for their
sensitive and reproducible detection. However, recent studies have revealed an alarming degree
of non-specific binding found in these antibodies, and whether the affinity meets the required
detection sensitivity is unclear. To address these challenges, we investigated whether directed
evolution could be applied to improve the affinity of a high-specific antibody targeting phospho-
threonine 231 of the human microtubule-associated protein tau. We developed a novel approach
using yeast surface display to quantify the specificity of PTM-specific antibodies. This approach
enabled us to measure its affinity and reveal its cross-reactivity towards the non-targets when we
affinity-matured the single-chain variable antibody fragment through directed evolution. We
demonstrate that directed evolution is a viable approach for obtaining high-affinity PTM-specific
antibodies, and highlight the importance of assessing the specificity in the antibody engineering
process.
The yeast surface display approach allows specificity measurement when the sequence of the
antibody is known, which makes it useful for screening antibody variants. However, for many
antibody users, the assay is not readily applicable for measuring specificity, since the antibody
sequence information may be unavailable. Therefore, we developed another assay that enables
PTM-specificity measurement for existing, commercially available antibodies. The assay relies
Dan Li – University of Connecticut, 2019
on multi-color flow cytometry in human embryonic kidney (HEK) cells. We defined a specificity
parameter termed Φ , which measures the fraction of non-specific signal in PTM-specific
antibodies, and successfully applied the HEK cell-based assay to measure the Φ values. We
validated the assay using antibodies with known specificity profiles, and used it to measure the
specificity of 7 widely used phospho-tau antibodies (AT270, AT8, AT100, AT180, PHF-6, TG-3,
and PHF-1) among others. We anticipate that the quantitative approach and parameter introduced
here will be widely adopted as a standard for reporting the specificity for phospho-tau antibodies,
and potentially for PTM-specific antibodies in general.
Engineering Affinity and Specificity of Antibodies Targeting Protein Modifications
Dan Li
B.S., Tianjin University, China, 2014
A Dissertation
Submitted in Partial Fulfillment of the
Requirements for the Degree of
Doctor of Philosophy
at the
University of Connecticut
2019
ii
Copyright by
Dan Li
2019
iii
Doctor of Philosophy Dissertation
Engineering Affinity and Specificity of Antibodies Targeting Protein Modifications
Presented by
Dan Li, B.S.
Major Advisor _________________________________________________________________
Dr. Yongku Cho
Associate Advisor ______________________________________________________________
Dr. Guoan Zheng
Associate Advisor ______________________________________________________________
Dr. Xudong Yao
Associate Advisor ______________________________________________________________
Dr. Yu Lei
University of Connecticut
2019
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First of all, I would like to thank my major advisor, Dr. Yongku Cho for the opportunity to learn
and study in his lab at UConn. The amount of patience and effort he was able to exhibit with
regard to my study and research is astounding. Our weekly discussions were always productive
and continually pushed me in the right direction. I thank him for helping my writing and
presentation skill, as well as my professional thinking. The work in this dissertation would not
have been possible without his guidance and support. I am so proud to have the opportunity to
work for him, be mentored by him, and I will look back on our work together fondly.
I would like to thank my associate advisors: Dr. Guoan Zheng, Dr. Xudong Yao and Dr. Yu Lei,
for their time and valuable guidance on my dissertation. I am also very thankful to Dr. Xudong
Yao and Dr. Benjamin Wolozin for their scientific input, kindness and time they dedicated to our
collaboration and research.
I am sincerely grateful to have worked with my lab mates: Shiyao Wang, Azady Pirhanov and
Monika Arbaciauskaite. It has been wonderful to see you and work with you in Dr. Cho’s lab. I
thank you all for both your professional and emotional support in the past. I also want to thank
all of my friends, both at UConn and elsewhere who made my life here pleasant and enjoyable.
In the end, I would like to acknowledge my family: my mom and my brother. Thank you for the
amount of love, encouragement, and support for my study and life. Without their unconditional
support, I could not have the chance to get here, to be the person I am today! I can’t express how
much I miss them at this moment! This work is dedicated to my beloved family.
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Table of Contents
APPROVAL PAGE ..................................................................................................................... iii
ACKNOWLEDGEMENTS ........................................................................................................ iv
List of tables.................................................................................................................................. ix
List of figures ................................................................................................................................. x
Chapter 1 Introduction ........................................................................................................... 1
Overview ............................................................................................................................. 1
Research objectives ............................................................................................................. 3
Significance and contributions ............................................................................................ 5
Organization of the thesis ................................................................................................... 7
References ........................................................................................................................... 8
Chapter 2 Background and literature review .................................................................... 10
Antibodies targeting protein PTMs ................................................................................... 10
2.1.1 Protein PTMs .................................................................................................... 10
2.1.2 PTM-specific antibodies ................................................................................... 11
Tau protein ........................................................................................................................ 12
2.2.1 Tau PTMs and Alzheimer’s Disease ................................................................. 14
2.2.2 Phosphorylation of tau ...................................................................................... 15
2.2.3 Commonly used phospho-tau antibodies .......................................................... 16
2.2.4 Specificity validation approaches of phospho-tau antibodies ........................... 18
Yeast surface display platform .......................................................................................... 18
References ......................................................................................................................... 21
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Chapter 3 Directed evolution of a picomolar-affinity, high-specificity antibody targeting
phosphorylated tau ..................................................................................................................... 26
Introduction ....................................................................................................................... 26
Materials and Methods ...................................................................................................... 29
3.2.1 Yeast surface display of pT231 scFv ................................................................ 29
3.2.2 Peptide modification and purification ............................................................... 30
3.2.3 Directed evolution of pT231 scFv .................................................................... 31
3.2.4 Purification of pT231 scFvs .............................................................................. 33
3.2.5 Fluorescent tagging of purified pT231 scFvs ................................................... 34
3.2.6 Immunocytochemistry in HEK 293 cells .......................................................... 34
3.2.7 Immunohistochemistry in brain tissue sections ................................................ 35
Results ............................................................................................................................... 36
3.3.1 Yeast surface display of a phospho-tau scFv .................................................... 36
3.3.2 Chemical labeling of peptides to generate distinctively labeled epitopes ........ 37
3.3.3 Verification of binding activity of yeast displayed pT231 scFv ....................... 38
3.3.4 Assessment of antibody phospho-specificity using yeast surface display ........ 40
3.3.5 Directed evolution of pT231 scFv for improved affinity .................................. 42
3.3.6 Characterization of affinity and specificity of scFv mutants ............................ 45
3.3.7 Cell and tissue section labeling using purified scFvs ....................................... 48
Discussion ......................................................................................................................... 52
Conclusion ........................................................................................................................ 55
Reference .......................................................................................................................... 57
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Chapter 4 High specificity of widely used phospho-tau antibodies validated using a
quantitative whole-cell based assay ........................................................................................... 62
Introduction ....................................................................................................................... 62
Materials and Methods ...................................................................................................... 64
4.2.1 Cloning of recombinant constructs of tau and mutant tau ................................ 64
4.2.2 Cell culture and transfection ............................................................................. 66
4.2.3 Tau antibodies ................................................................................................... 66
4.2.4 Quantitation of antibody specificity .................................................................. 68
4.2.5 Measurement of Φ: validation of phospho-tau antibody specificity ................ 69
4.2.6 Immunocytochemistry in HEK293FT cells ...................................................... 71
4.2.7 Mouse hippocampus dissection ........................................................................ 71
4.2.8 HEK293FT and mouse primary cell lysate preparation and western blotting .. 72
Results ............................................................................................................................... 73
4.3.1 Φ: A generalizable parameter for measuring the specificity of phospho-tau
antibodies .......................................................................................................................... 73
4.3.2 Co-expression of glycogen synthase kinase-3b enhances tau phosphorylation 77
4.3.3 Quantifying the phospho-site specificity of commonly used phospho-tau
antibodies .......................................................................................................................... 78
4.3.4 Non-specific binding to cells in phospho-tau antibodies .................................. 81
Discussion ......................................................................................................................... 84
Conclusion ........................................................................................................................ 88
References ......................................................................................................................... 90
Chapter 5 Conclusion and future perspectives .................................................................. 93
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Conclusion ........................................................................................................................ 93
Future perspectives ........................................................................................................... 93
5.2.1 Avidity effect on antibody screening assay ...................................................... 93
5.2.2 Application of binding proteins in optogenetics ............................................... 97
Reference ........................................................................................................................ 112
ix
Table 4.1 Primers for Golden Gate assembly of plasmids ............................................................ 65
Table 4.2 Primers used for site-directed mutagenesis .................................................................. 65
Table 4.3 Phospho-tau antibodies ................................................................................................. 67
x
Figure 2.1 Antibody-phosphopeptide interaction revealed by X- ray crystallography ................ 12
Figure 2.2 The transcription, alternative splicing and PTMs of tau. ............................................ 13
Figure 2.3 Putative phosphorylation sites on tau protein and epitopes specific for major tau
antibodies ...................................................................................................................................... 16
Figure 2.4 A schematic overview of scFv scaffold displayed on the surface of yeast ................. 20
Figure 3.1 Schematic representation of antibody specificity quantification using yeast surface
display ........................................................................................................................................... 37
Figure 3.2. Modified peptides purification and confirmation ....................................................... 38
Figure 3.3 Binding activity of yeast displayed pT231 scFv ......................................................... 39
Figure 3.4 Assessment of CDR mutations on phosphopeptide binding of yeast displayed pT231
scFv ............................................................................................................................................... 40
Figure 3.5 Assessment of the specificity of yeast displayed pT231 scFv .................................... 41
Figure 3.6 Directed evolution of yeast displayed pT231 scFv variants with improved affinity .. 43
Figure 3.7 Amino acid sequence of individual mutants isolated from the screen for improved
affinity ........................................................................................................................................... 44
Figure 3.8 High occurrence mutations identified from the screen mapped on WT pT231 scFv
structure ......................................................................................................................................... 45
Figure 3.9 Characterization of affinity and specificity of individual scFvs ................................. 47
xi
Figure 3.10 Confocal microscopy images of HEK293FT cell labeled using purified pT231 scFvs
....................................................................................................................................................... 49
Figure 3.11 Quantification of cell labeling fluorescence intensity using purified pT231 scFvs .. 50
Figure 3.12 Tissue labeling using purified pT231 scFvs .............................................................. 51
Figure 4.1 A ratiometric assay to quantify the specificity of phospho-tau antibodies. ................ 74
Figure 4.2 Validation of the assay using well characterized anti-tau antibodies .......................... 76
Figure 4.3 Measurement of Φala and ΦλPP of phospho-tau antibodies. ..................................... 77
Figure 4.4 Co-expression of GSK-3b in HEK293FT cells enhances phosphorylation of tau. ..... 78
Figure 4.5 Specificity measurements for commonly used phospho-tau antibodies ...................... 81
Figure 4.6 Flow cytometry plots showing binding of isotype-matching control antibodies under
the specificity quantification assay conditions ............................................................................. 82
Figure 4.7 Non-specific binding to HEK293FT cells in phospho-tau antibodies ......................... 84
Figure 5.1 Schematic of the Streptavidin-biotin interaction ......................................................... 96
Figure 5.2 Schematic of yeast surface display formats ................................................................. 97
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Overview
Antibodies that target post-translationally modified proteins are widely used in biology and
clinical investigations. Antibodies exist that can recognize a single modified amino acid (e.g.,
phosphorylated tyrosine) within the context of diverse nearby amino acid sequence (1,2).
However, most widely used are antibodies binding to a modification site in a defined target
protein sequence, herein referred to as post-translational modification (PTM)-specific antibodies.
Due to the large number of experimentally identified post-translational modification sites (>
87,000) (3), the demand for PTM-specific antibodies continue to grow. Due to the dynamic and
heterogeneous nature of PTMs, the affinity and specificity of PTM-specific (4) antibodies are
critical for sensitive and reproducible detection of PTMs (5). Only a fraction of a protein is
modified at any given time, and the dynamics may vary within a tissue across different cell types
(6-8). Therefore, PTM-specific antibodies should interact only with the modified target site, with
high affinity to capture potentially a small fraction of the modified form.
However, generating high affinity antibodies with high PTM-specificity remains to be
challenging. Existing methods for generating PTM-specific antibodies rely on immunization
approaches, but often fail or produce low affinity antibodies. This is due to the fact that the
peptide epitopes containing PTMs are weak immunogens in mice (9), PTMs undergo rapid
degradation in vivo regardless of the immunization route (10), and that B-cell response shows an
affinity ceiling (11,12). Therefore, screening recombinant antibody libraries in vitro is thought to
be a promising alternative. However, in vitro library screening strategies for PTM-specific
antibodies also tend to result in low affinity antibodies (Kd ~ 100 nM to µM range) (13-15). This
trend has been observed for screening phospho-specific antibodies, but it extends to antibodies
2
targeting other PTMs, such as sulfotyrosine (16). The specificity of PTM-specific antibodies is
recently gaining attention as a critical factor. In case of the microtubule-associated protein tau,
although phospho-specific antibodies are highly sought after for many target proteins, often at
multiple phosphorylation sites, they are currently available in limited cases, and often suffer
from poor specificity and low affinity (17-19).
High affinity and specificity antibodies against phosphorylated tau will enable the detection in
low concentration samples. This would be a major factor in developing diagnostic assays using
patient samples such as cerebrospinal fluid (CSF). Recent studies show that majority of existing
monoclonal antibodies cannot detect phosphorylated tau, due to low affinity. Affinity of
antibodies that recognize PTM tau is also an important factor in developing passive
immunotherapy strategies. Studies in mouse models have shown that for certain tau epitopes,
targeting antibody affinity is not a major factor, while for antibodies targeting soluble tau affinity
was highly correlated with efficacy (20). High affinity antibodies against phosphorylated tau will
be necessary for systematic studies to determine key factors to efficacy.
This work focuses on the development of assays for generating and characterizing PTM-specific
antibodies with high specificity and affinity. To assess the specificity of PTM-targeting
antibodies, we develop an experimental approach based on yeast surface display to quantify
single-chain variable fragment (scFv) binding against a defined set of structurally similar
epitopes. We generate a picomolar affinity scFv targeting phosphorylated T231 of human tau
using directed evolution. The yeast surface display approach allows specificity measurement
when the sequence of the antibody is known, which makes it useful for screening antibody
variants. However, for many antibody users, the assay is not readily applicable for measuring
specificity, since the antibody sequence information may be unavailable. Therefore, we report a
3
robust flow cytometry assay using human embryonic kidney (HEK) cells that enables the
determination of a specificity parameter termed Φ, which measures the fraction of non-specific
signal in antibody binding. We apply our assay to measure the specificity of 7 widely used
phospho-tau antibodies (AT270, AT8, AT100, AT180, PHF-6, TG-3, and PHF-1) among others.
Research objectives
It is the aim of this work to engineer and assess affinity and specificity of antibodies targeting
protein PTM, particularly in tau phosphorylation. The objective of this research can be broken
down into 2 aims:
(1) Develop a screening assay to generate high-affinity and specific PTM-specific antibody using
yeast surface display. According to the x-ray crystal structural of antibody-phosphoepitope
complex, it’s plausible that high-affinity phospho-specific antibodies selectively stabilize either
the peptide sequence or phosphate interactions, leading to increase in cross reactivity towards the
non-phosphoepitope or irrelevant phosphorylation sites. Therefore, we hypothesized that a trade-
off between affinity and specificity might exist in PTM-specific antibodies. To test this
hypothesis, we sought to assess the impact of in vitro affinity maturation on the specificity of
PTM-targeting antibodies.
The specific aims of this project were:
i. Development of an assay to quantify specificity of phospho-specific antibody using
yeast surface display
• Yeast surface display of a phospho-tau-specific scFv
• Chemically labelling peptides to generate distinctively labeled epitopes
• Assessment of scFv phospho-specificity using yeast surface display
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ii. Engineering antibody phospho-specificity and affinity through directed evolution
• Creation of a yeast displayed scFv library using random mutagenesis and DNA shuffling
• Affinity maturation of yeast displayed scFvs using FACS
iii. Characterization of affinity and specific of scFv mutants
• Validation of scFv mutants using yeast surface display
• Testing the good performing scFv mutants in cell and tissue section labeling
In summary, the outcomes of this project were: introducing a yeast surface display approach to
quantify the specificity of PTM-specific antibodies, and developed a screening assay based on it
to generate PTM-specific antibodies with improved affinity and high specificity through directed
evolution.
(2) Introduce a quantitative whole-cell based assay to validate PTM-specific antibody specificity.
Antibodies binding to phosphorylated tau (phospho-tau) are extensively used in research and
being adopted in clinical diagnosis. However, few validation data have been published and
existing specificity characterization methods relied on immunoblotting approaches which are not
ideal for quantitative measurement due to frequent signal saturation in immunoblotting
experiments. To fill this critical need, we introduced a parameter termed Φ that quantifies
specificity of phospho-tau antibodies and established a generalizable, robust assay to measure it.
The specific aims of this project were:
i. Development of a whole-cell based approach to quantify phospho-tau antibody
specificity using flow cytometry
• HEK293 cells expressing fluorescent tagged full-length tau
• Validation of the assay using previously characterized anti-tau antibodies
• Defining a parameter Φ to quantitatively measure the specificity phospho-tau antibodies
5
ii. Quantifying the phospho-specificity of commonly used phospho-tau antibodies
• Measurement of specificity Φ of commonly used phospho-tau antibodies using flow
cytometry
• Assessment of non-specific binding in phospho-tau antibodies by performing cell
labeling and western blotting
In summary, the outcomes of this project were: providing a whole-cell based approach to
validate the specificity of phospho-tau antibodies, and determined a parameter Φ for phospho-tau
antibodies based on it, that enable quantitative comparison of specificity. We anticipate that this
parameter will be widely adopted in guiding users of the specificity of phospho-tau antibodies,
and potentially PTM-specific antibodies in general.
Significance and contributions
High quality antibodies require high specificity and affinity toward the targets. The specificity
and affinity of PTM-specific antibodies are critical for sensitive and reproducible detection of
protein PTM. However, the current methods for generating PTM-specific antibodies have
drawbacks that make is hard to identify high quality antibodies; and many PTM-specific
antibodies have low affinity and poor specificity. This work improved the success rate of
generating high quality PTM-specific antibodies, and provided an assay to quantitatively
measure the specificity of those antibodies.
This work allows for contributions in the following fields:
i. Introduced a yeast surface display approach to quantify antibody specificity
ii. Developed a screening assay to generate antibodies with improved affinity and high
specificity using yeast surface display
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iii. Generated a pico-molar affinity scFv targeting phosphorylated T231 of human tau using
directed evolution
iv. Provided information on structure analysis of antibody-antigen interaction
v. Enabled small binders in cell and tissue staining
vi. Developed a whole-call based approach to quantitatively quantify the specificity of PTM-
specific antibodies
vii. Introduced a parameter Φ to measure the specificity of PTM-specific antibodies, which
will be widely adopted in guiding the use PTM-specific antibodies in general
Manuscripts and publications have been resulted from this work as listed below:
1. Li, D., Cho, Y.K.*, High specificity of widely used phospho-tau antibodies validated
using a quantitative whole-cell based assay, in revision, 2019
2. Li, D., Wang, L., Maziuk, B.F., Yao, X., Wolozin, B., Cho, Y.K.*, Directed evolution of
a picomolar affinity high-specificity antibody targeting phosphorylated tau, Journal of
Biological Chemistry, 2018, 293, 12081-12094
3. Knowlton, S., Li, D., Ersoy, F., Cho, Y.K.*, Tasoglu S.*, Building Blocks for Bottom-
Up Neural Tissue Engineering: Tools for In Vitro Assembly and Interrogation of Neural
Circuits (book chapter), Neural Engineering, 2016
4. Cho, Y.K.*, Li, D., Optogenetics: Basic Concepts and Their Development (book chapter),
Methods in Molecular Biology, 2016
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Organization of the thesis
Chapter 2 provides an overview of the background, concepts and significances of PTM-specific
antibodies. It begins with a brief introduction of protein PTMs and the antibodies that targets
such antigens. Antibodies targeting tau PTMs are highlighted, specifically discussing the
antibodies targeting phosphorylated tau that is associated with Alzheimer’s Disease. In the end of
this chapter, we review a yeast surface display technique, that is vital in in vitro recombinant
antibody screening assays.
In Chapter 3, we introduce an approach using yeast surface display to quantify the specificity of
phospho-specific antibodies. We develop a screening assay based on it to generate PTM-specific
antibodies with improved affinity and high specificity. We demonstrate that in vitro affinity
maturation leads to cross-reactivity in PTM-specific antibodies. However, quantifying the
specificity allows identifying an antibody with improved affinity and specificity.
In Chapter 4, we provide a whole-cell based approach to validate the specificity of phospho-tau
antibodies. We introduce the parameter Φ for phospho-tau antibodies, that enable quantitative
comparison of specificity. Using our assay, we quantified the specificity Φ of 7 widely used
phospho-tau antibodies (AT270, AT8, AT100, AT180, PHF-6, TG-3, and PHF-1) as well as 2
additional phospho-tau antibodies (1H6L6 and pS404).
Chapter 5 presents a summary of this work and provides the future perspectives of this work.
First, we will assess the avidity effect of the yeast surface displayed scFvs for the targets, and
apply it in developing a novel in vitro screening assay. Second, we will develop small binding
protein for tau, combined with optogenetics tools allows the visualization of complex structures
such as synapse.
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References
1. Rush, J., Moritz, A., Lee, K. A., Guo, A., Goss, V. L., Spek, E. J., Zhang, H., Zha, X. M., Polakiewicz, R. D., and Comb, M. J. (2005) Immunoaffinity profiling of tyrosine phosphorylation in cancer cells. Nature biotechnology 23, 94-101
2. van der Mijn, J. C., Labots, M., Piersma, S. R., Pham, T. V., Knol, J. C., Broxterman, H. J., Verheul, H. M., and Jimenez, C. R. (2015) Evaluation of different phospho-tyrosine antibodies for label-free phosphoproteomics. J Proteomics 127, 259-263
3. Khoury, G. A., Baliban, R. C., and Floudas, C. A. (2011) Proteome-wide post-translational modification statistics: frequency analysis and curation of the swiss-prot database. Sci Rep 1
4. Min, S. W., Chen, X., Tracy, T. E., Li, Y., Zhou, Y., Wang, C., Shirakawa, K., Minami, S. S., Defensor, E., Mok, S. A., Sohn, P. D., Schilling, B., Cong, X., Ellerby, L., Gibson, B. W., Johnson, J., Krogan, N., Shamloo, M., Gestwicki, J., Masliah, E., Verdin, E., and Gan, L. (2015) Critical role of acetylation in tau-mediated neurodegeneration and cognitive deficits. Nature medicine
5. Lothrop, A. P., Torres, M. P., and Fuchs, S. M. (2013) Deciphering post-translational modification codes. FEBS Lett 587, 1247-1257
6. Bayes, A., and Grant, S. G. (2009) Neuroproteomics: understanding the molecular organization and complexity of the brain. Nat Rev Neurosci 10, 635-646
7. Prabakaran, S., Lippens, G., Steen, H., and Gunawardena, J. (2012) Post-translational modification: nature's escape from genetic imprisonment and the basis for dynamic information encoding. Wiley Interdiscip Rev Syst Biol Med 4, 565-583
8. Witze, E. S., Old, W. M., Resing, K. A., and Ahn, N. G. (2007) Mapping protein post-translational modifications with mass spectrometry. Nature methods 4, 798-806
9. Brumbaugh, K., Johnson, W., Liao, W. C., Lin, M. S., Houchins, J. P., Cooper, J., Stoesz, S., and Campos-Gonzalez, R. (2011) Overview of the generation, validation, and application of phosphosite-specific antibodies. Methods in molecular biology 717, 3-43
10. Czernik, A. J., Girault, J. A., Nairn, A. C., Chen, J., Snyder, G., Kebabian, J., and Greengard, P. (1991) Production of phosphorylation state-specific antibodies. Methods in enzymology 201, 264-283
11. Batista, F. D., and Neuberger, M. S. (1998) Affinity dependence of the B cell response to antigen: a threshold, a ceiling, and the importance of off-rate. Immunity 8, 751-759
12. Foote, J., and Eisen, H. N. (1995) Kinetic and affinity limits on antibodies produced during immune responses. Proceedings of the National Academy of Sciences of the United States of America 92, 1254-1256
13. Koerber, J. T., Thomsen, N. D., Hannigan, B. T., Degrado, W. F., and Wells, J. A. (2013) Nature-inspired design of motif-specific antibody scaffolds. Nature biotechnology 31, 916-921
14. Shih, H. H., Tu, C., Cao, W., Klein, A., Ramsey, R., Fennell, B. J., Lambert, M., Ni Shuilleabhain, D., Autin, B., Kouranova, E., Laxmanan, S., Braithwaite, S., Wu, L., Ait-Zahra, M., Milici, A. J., Dumin, J. A., LaVallie, E. R., Arai, M., Corcoran, C., Paulsen, J. E., Gill, D., Cunningham, O., Bard, J., Mosyak, L., and Finlay, W. J. (2012) An ultra-
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specific avian antibody to phosphorylated tau protein reveals a unique mechanism for phosphoepitope recognition. The Journal of biological chemistry 287, 44425-44434
15. Feldhaus, M. J., Siegel, R. W., Opresko, L. K., Coleman, J. R., Feldhaus, J. M., Yeung, Y. A., Cochran, J. R., Heinzelman, P., Colby, D., Swers, J., Graff, C., Wiley, H. S., and Wittrup, K. D. (2003) Flow-cytometric isolation of human antibodies from a nonimmune Saccharomyces cerevisiae surface display library. Nature biotechnology 21, 163-170
16. Kehoe, J. W., Velappan, N., Walbolt, M., Rasmussen, J., King, D., Lou, J., Knopp, K., Pavlik, P., Marks, J. D., Bertozzi, C. R., and Bradbury, A. R. (2006) Using phage display to select antibodies recognizing post-translational modifications independently of sequence context. Molecular & cellular proteomics : MCP 5, 2350-2363
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20. Congdon, E. E., Lin, Y., Rajamohamedsait, H. B., Shamir, D. B., Krishnaswamy, S., Rajamohamedsait, W. J., Rasool, S., Gonzalez, V., Levenga, J., Gu, J., Hoeffer, C., and Sigurdsson, E. M. (2016) Affinity of Tau antibodies for solubilized pathological Tau species but not their immunogen or insoluble Tau aggregates predicts in vivo and ex vivo efficacy. Mol Neurodegener 11, 62
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Antibodies targeting protein PTMs
Antibodies that target proteins post-translation modifications (PTMs) are widely used in biology
and clinical investigations. Some antibodies can bind to a single modified amino acid (e.g.,
phosphorylated tyrosine) within the context of diverse nearby peptide sequences (1,2). However,
most widely used are antibodies specific to a modification site in a defined target protein
sequence, herein referred to as PTM-specific antibodies. Due to a large number of
experimentally identified post-translational modification sites (> 87,000) (3), the demand for
PTM-specific antibodies continues to grow.
2.1.1 Protein PTMs
In the last few decades, researchers discovered that the human proteome is more complex than
human genome. The increase of the complexity of proteome to genome is facilitated by many
different mechanisms, including PTMs. PTM is covalent and enzymatic modification of proteins
following protein biosynthesis. It occurs after the mRNA is translated into the protein sequence
in ribosomes. PTMs process the protein by adding different biochemical group (such as
phosphorylation, methylation), or adding different small protein molecule (such as
ubiquitination), or splicing protein sequence at specific sites. PTM is involved in nearly all
aspect of biological, and plays important role in regulating protein level and function. For
instance, phosphorylation, which is the most common PTM, alters biological activity of enzymes
(3). Many enzymes and receptors are switched "on" or "off" by phosphorylation and
dephosphorylation.
PTM is a key mechanism that enables reversible regulation of protein activity, conformation,
localization, turnover rate, and interactions. Abnormal PTMs result in dysregulation of these
11
processes and are often associated with pathological conditions, including heart disease, cancer,
and neurodegeneration (4-8). In case of tau protein, numerous abnormal PTMs regulate its
biology, and are associated with several neurodegenerative diseases including Alzheimer’s
Disease (AD), collectively referred to as tauopathies (9).
2.1.2 PTM-specific antibodies
Since protein PTMs are included in many pathological conditions, detection of protein PTMs is
valuable in identifying the cause and tracking the progression of these diseases. However, robust
detection of PTMs is challenging due to their dynamic and heterogeneous nature. Only a fraction
of a protein is modified at any given time, and the dynamics may vary within a tissue across
different cell types (10-12). Therefore, the affinity and specificity of PTM-specific antibodies are
critical for sensitive and reproducible detection of PTMs (13). PTM-specific antibodies should
interact only with the modified target site, with high affinity to capture potentially a small
fraction of the modified form.
Recently, an x-ray crystal structure of an antibody in complex with a phosphoepitope peptide of
tau (PDB code 4GLR) revealed a recognition motif, in which a sector of residues from the
complementarity determining region (CDR)-H2 interact with the phosphate group of
phosphorylated threonine 231of tau and another sets of CDR residues contacting the peptide
sequence (Figure 2.1) (14). Such distinct recognition sectors were also found in structural
analyses of other phospho-specific antibodies (15).
12
Figure 2.1 Antibody-phosphopeptide interaction revealed by X- ray crystallography The interaction is mediated by recognition of the phosphate group by CDR-H2 (solid) and the surrounding residues by other CDRs(dashed). Image generated based on PDB code 4GLR.
In case of tau PTM-specific antibodies, their affinity and specificity are particularly important,
since they dictate their efficacy in a wide range of applications. High-affinity antibodies are
superior in detecting tau PTMs from clinical samples (16), but mouse model experiments suggest
that high specificity antibodies may be more beneficial for passive immunotherapy targeting tau
(17).
Tau protein
Tau is a microtubule-associated protein tau (MAPT) which are found mostly in neurons. Its
major functions are to promote microtubule assembly and maintain the stability of the neurons
through binding to the microtubules. Tau has two ways to control microtubule stability: tau
isoforms and tau phosphorylation.
13
Human MAPT gene for encoding tau protein is located on the human chromosome 17q21,
having 16 exons (18). Alternative splicing of exons 2,3 and 10 leads to formation of six tau
isoforms which are composed of 352 - 441 amino acid residues with molecular weight of ~ 37 -
46 kDa (Figure 2.2) (19,20). Tau isoforms contain either 0, 1 or 2 inserts of 29 amino acids at
the N-terminus (exon 2 and 3), and 3 or 4 microtubule-binding repeats at the C-terminus (exon
10). Six isoforms of tau have been found in adult human brains (21).
Figure 2.2 The transcription, alternative splicing and PTMs of tau. By transcription and alternative splicing at exons 2, 3 and 10 of tau mRNA, six isoforms of tau protein are generated from a single gene located on the chromosome 17q21 in human. Different tau isoforms are named according to the numbers of the N-terminal inserts and/or C-terminal microtubule-binding domain (such as 2N4R) or according to the clones (such as Tau40) (19).
14
2.2.1 Tau PTMs and Alzheimer’s Disease
The complexity of the tau isoforms can be further increased by PTMs (Figure 2.2). During
normal development, tau protein undergoes various PTM, including phosphorylation,
glycosylation, acetylation, ubiquitination, truncation etc. Recent studies suggest that the PTMs of
tau may participate in the regulations of intracellular signal transduction, development and
viability of the neurons (22).
The abnormal PTM, such as hyper-phosphorylation, has been found in a number of
neurodegenerative disorder, known as tauopathy, such as AD, corticobasal degeneration, Down’s
syndrome. Phosphorylation sites at or near the repeat elements of the microtubule binding
domains are particularly important for the pathophysiology of tauopathy (23). An increase in tau
phosphorylation, on average from 2 - 3 to 7 - 8 phosphate per molecule (23) results in
preferential phosphorylation at sites that decrease microtubule binding (24,25), and increases its
propensity to form oligomers and high order aggregates (23,26). Levels of tau oligomers showed
a strong correlation with memory loss in a mouse model of tauopathy (27). Acetylation at
specific lysine residues interferes with ubiquitination and subsequent degradation by the
proteasome, resulting in accumulation of phosphorylated tau and associated toxicity in mouse
models (28-31).
The tau hypothesis states that, in AD brains, tau becomes hyperphosphorylated, resulting in
reduced affinity for microtubules and self-aggregating into paired helical filament (PHF), leading
to formation of neurofibrillary tangles (32,33). This process causes the microtubules
depolymerisation, and finally breaks the neurons. Therefore, these phosphorylated neurofibrillary
tangles are often seen in AD brain and thought to be a hallmark of AD.
15
AD is a progressive neurodegenerative disease, it’s the most well-known form of dementia,
accounting for 50-60% of all cases (34). AD patients often suffer from memory loss, confusion
and behavioral issues. Gradually, bodily functions are lost, ultimately leading to death. In 2018,
50 million people have dementia in worldwide, and there will be one new case every 3 second,
so that more than 150 million people will live with dementia by 2050. In 2018 the global societal
cost of dementia is 1 trillion US dollars, and it will rise to 2 trillion by 2030. Until now, there are
no drugs available that can prevent or even slower this course.
2.2.2 Phosphorylation of tau
On the basis of tau hypothesis, abnormal PTM of tau is currently the most common disease-
modifying target. Among these PTMs, tau phosphorylation is the most common and the best-
studied. In AD brains, the level of the hyper-phosphorylated tau is 3-4 fold higher than that of
tau isolated from normal brains (35,36). There are 85 serine, threonine, or tyrosine
phosphorylation sites in the longest human brain tau isoform, and 45 sites have been identified in
AD brain samples by using mass spectra or antibodies recognition (Figure 2.3) (37,38). And
they are clustered in the proline-rich domain (PRD) and C-terminal region of tau.
16
Figure 2.3 Putative phosphorylation sites on tau protein and epitopes specific for major tau antibodies Red color denotes amino acids phosphorylated in AD brain, green in both AD and normal brain, blue in normal brain, while black color means that those phosphorylation sites have not been fully characterized yet. Tau antibodies specific for phospho-tau epitopes are given in purple, while pink color denotes antibodies specific for non-phosphorylated tau epitopes (38).
2.2.3 Commonly used phospho-tau antibodies
Antibodies targeting tau phosphorylation (phospho-tau) are extensively used in research and
being adopted in clinical diagnosis (39-44). According to antibody databases (Antibodypedia and
Alzforum), thousands of anti-tau antibodies are currently available, including >180 unique
phospho-tau antibodies. Epitopes recognized by some commonly used phospho-tau and
nonphospho-tau antibodies are shown in Figure 2.3 (38).
17
AT270. This antibody is against phosphorylated epitope T181(45,46), which is in PRD of tau.
AT270 is produced using PHF tau as the immunogen, and it does not recognize recombinant tau,
but does so after phosphorylation with brain extract.
AT8. This antibody is against phosphorylation epitope S199 and/or S202 and/or T205 (45,46),
which are in PRD of tau. Previous study has shown that AT8 recognition requires only S202 to
be phosphorylated in tau (47). AT8 recognizes all isoforms of PHF tau but not tau from normal
mammalian brain, thus the antibody is specific for the Alzheimer state and has the advantage of
reporting on the state of phosphorylation (48).
AT100. This antibody is against doubly phosphorylated epitope T212/S214 (49-52), which are in
PRD of tau. AT100 epitope is formed only when a specific order of phosphorylation occurs (53).
It has also been shown that Thr212 and Ser214 are sequentially phosphorylated by glycogen
synthase kinase 3β (GSK-3b) and protein kinase A (PKA), respectively (53).
AT180. This antibody is against phosphorylated epitope T231 (45,51,54), which is in PRD of tau.
AT180 is produced using PHF tau as the immunogen, and it does not recognize recombinant tau,
but does so after phosphorylation with brain extract.
PHF6. This antibody is against phosphorylated epitope T231 (55,56), which is in PRD of tau.
PHF6 recognizes PHF tau containing phosphorylated T231.
TG-3. This antibody is sensitive to the conformation changes of phosphorylated Thr231, which
is in PRD of tau, it recognizes neurofibrillary tangles of tau in AD, while not reacts to normal tau
(51,57,58).
PHF1. This antibody is against doubly phosphorylated epitope S396/S404 (50,59,60), which are
in C-terminal region of tau. PHF1 recognizes PHF tau containing either individually
18
phosphorylated S396 or S404, but is more sensitive in detection when both serines are
phosphorylated.
2.2.4 Specificity validation approaches of phospho-tau antibodies
Recent studies have shown that some phospho-tau antibodies lack specificity (61-63). Existing
specificity characterization methods relied on immunoblotting approaches using either synthetic
peptides, cell lysates or extracted PHF tau (61-63). Synthetic peptides allow binding
measurement to exactly defined phosphorylation sites, but is prohibitively costly to cover all
phospho-sites across the full-length tau. Cell lysates allow detection of non-specific binding to
non-tau proteins (61), but cannot measure cross-reactivity to non-modified tau or other
phosphorylation sites. Extracted PHF-tau from AD brain homogenates allows characterizing
antibodies that do not recognize normal human adult tau and measuring the cross-reactivity to
nonphospho-tau by treatment with phosphatase, but cannot capture the phospho-site specificity
(63). Steinhilb and colleagues have generated transgenic flies expressing tau with alanine point
mutations at several serine-proline and threonine-proline motifs of human tau (64). In this study,
they used fly head extracts from these transgenic lines to perform western blotting using
phospho-tau antibodies. In principle, this approach may be applied for validating phospho-tau
antibodies. However, all of these approaches are not ideal for quantitative measurement due to
frequent signal saturation in immunoblotting experiments (65).
Yeast surface display platform
Yeast surface display is a powerful method for isolating and engineering antibodies to increase
their affinity and specificity (66). Yeast surface display has been used to engineer antibodies
targeting several proteins, including T cell receptors (67), botulinum neurotoxin (68), and
19
huntingtin protein (69). Yeast display libraries, which has 107-109 transformants, are smaller than
phage or ribosome display libraries, which is 1011-1013, but have many advantages of eukaryotic
expressed proteins. They have eukaryotic expression, processing, and secretory pathway, which
suggest more complex PTMs of proteins (70).
In the yeast surface display system (Fig. 2.4), the antibody is displayed as a single chain variable
fragment (scFv) in which the heavy and light variable chains are connected by a flexible
polypeptide linker (Gly4Ser)3. The scFv is fused to the C-terminal end of agglutinin Aga2 protein,
which forms two disulfide bonds to the agglutinin Aga1 protein on the yeast surface. Both Aga1
and Aga2 protein are controlled under a galactose-inducible promoter. Each yeast cell displays
40,000-50,000 scFvs (66,71), and can be detected through immunofluorescence labeling of either
the c-myc or the hemagglutinin (HA) epitope tag flanking the scFv. The binding to an antigen of
interest can be detected by labeling the yeast cell with antigen and a secondary reagent
conjugated with fluorophore. Yeast surface display library enables quantitative screening using
fluorescence-activated cell sorting (FACS), and the antigen binding signal can be normalized for
expression, eliminating impacts due to host expression bias.
20
Figure 2.4 A schematic overview of scFv scaffold displayed on the surface of yeast The scFv is C-terminal fusion to the Aga2 and flanked by two epitopes: HA at the N-terminus and c-Myc at the C-terminus. The Aga2p forms two disulfide bonds with the Aga1p, which is anchored to the cell wall. Antigen binding is detected with a fluorophore FITC. Display of scFv is detected with a primary mouse anti-c-Myc antibody and a phycoerythrin-conjugated mouse secondary antibody (66).
Once the candidate scFvs are isolated from the library, they are engineered for improved affinity
and specificity (72) using random mutagenesis through error-prone PCR with nucleotide analogs
(73). The mutation frequency can be controlled by altering the number of PCR cycles and the
amount of nucleotide analogs. In addition, DNA shuffling (74,75) can be used to introduce
diversity into the library. The mutagenized library is created by transformation of yeast cells with
the mutated products and linearized vector through homologous recombination in yeast (75).
21
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66. Boder, E. T., and Wittrup, K. D. (1997) Yeast surface display for screening combinatorial polypeptide libraries. Nature biotechnology 15, 553-557
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67. Kieke, M. C., Cho, B. K., Boder, E. T., Kranz, D. M., and Wittrup, K. D. (1997) Isolation of anti-T cell receptor scFv mutants by yeast surface display. Protein Eng 10, 1303-1310
68. Razai, A., Garcia-Rodriguez, C., Lou, J., Geren, I. N., Forsyth, C. M., Robles, Y., Tsai, R., Smith, T. J., Smith, L. A., Siegel, R. W., Feldhaus, M., and Marks, J. D. (2005) Molecular evolution of antibody affinity for sensitive detection of botulinum neurotoxin type A. J Mol Biol 351, 158-169
69. Colby, D. W., Garg, P., Holden, T., Chao, G., Webster, J. M., Messer, A., Ingram, V. M., and Wittrup, K. D. (2004) Development of a human light chain variable domain (V(L)) intracellular antibody specific for the amino terminus of huntingtin via yeast surface display. J Mol Biol 342, 901-912
70. Bowley, D. R., Labrijn, A. F., Zwick, M. B., and Burton, D. R. (2007) Antigen selection from an HIV-1 immune antibody library displayed on yeast yields many novel antibodies compared to selection from the same library displayed on phage. Protein Eng Des Sel 20, 81-90
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26
Introduction
Antibodies that target post-translationally modified proteins are widely used in biology and
clinical investigations. Some antibodies can bind to a single modified amino acid (e.g.,
phosphorylated tyrosine) within the context of diverse nearby peptide sequences (1,2). However,
most widely used are antibodies specific to a modification site in a defined target protein
sequence, herein referred to as PTM-specific antibodies. Due to a large number of
experimentally identified post-translational modification sites (3), the demand for PTM-specific
antibodies continues to grow.
PTM is a key mechanism that enables reversible regulation of protein activity, conformation,
localization, turnover rate, and interactions. Abnormal PTMs result in dysregulation of these
processes and are often associated with pathological conditions. In case of the microtubule-
associated protein tau, numerous abnormal PTMs regulate its biology, and are associated with
several neurodegenerative diseases including AD, collectively referred to as tauopathies (4). Tau
contains 85 putative phosphorylation sites (5) and 14 putative acetylation sites (6).
Phosphorylation sites at or near the repeat elements of the microtubule binding domains are
particularly important for the pathophysiology of tauopathy (7). An increase in tau
phosphorylation, on average from 2-3 to 7-8 phosphate per molecule (7) results in preferential
phosphorylation at sites that decrease microtubule binding (8,9), and increases its propensity to
form oligomers and high order aggregates (7,10). Levels of tau oligomers showed a strong
correlation with memory loss in a mouse model of tauopathy (11). Acetylation at specific lysine
residues interferes with ubiquitination and subsequent degradation by the proteasome, resulting
27
in accumulation of phosphorylated tau and associated toxicity in mouse models (12-15).
Therefore, detection of tau PTMs at specific sites is valuable in identifying the cause and
tracking the progression of tauopathies.
However, robust detection of PTMs is challenging due to their dynamic and heterogeneous
nature. Only a fraction of a protein is modified at any given time, and the dynamics may vary
within a tissue across different cell types (16-18). Therefore, the affinity and specificity of PTM-
specific antibodies are critical for sensitive and reproducible detection of PTMs (19). PTM-
specific antibodies should interact only with the modified target site, with high affinity to capture
potentially a small fraction of the modified form. In case of tau PTM-specific antibodies, their
affinity and specificity are particularly important, since they dictate their efficacy in a wide range
of applications. High-affinity antibodies are superior in detecting tau PTMs from clinical
samples (20), but mouse model experiments suggest that high specificity antibodies may be more
beneficial for passive immunotherapy targeting tau (21).
Even though many PTM-specific antibodies exist, high-affinity antibodies with high PTM-
specificity are rarely found. Rodent immunization approaches often fail or produce low-affinity
antibodies. This is because the peptide epitopes containing PTMs are weak immunogens in mice
(22), PTMs undergo rapid degradation in vivo regardless of the immunization route (23), and B-
cell response shows an affinity ceiling (24,25). Screening recombinant antibody libraries in vitro
is a promising alternative. However, in vitro library screening strategies tend to generate PTM-
specific antibodies with a low or moderate affinity (Kd ~ µM to 100 nM range) (26-29).
Obtaining high specificity antibodies targeting PTMs is even more challenging. In fact, many
commercially available PTM-specific antibodies have not been validated for their specificity.
Recent evidences using various approaches, including immunoassays using synthetic peptides
28
and peptide arrays, have shown that indeed a significant fraction of existing PTM-targeting
antibodies lack specificity (30-34). These studies have identified major species that non-
specifically bind to the antibodies, including non-modified epitopes with the target amino acid
sequence and those with modifications at non-target sites (31,34).
Recently, X-ray crystal structures of phospho-specific antibodies have revealed a key insight on
how PTM-targeting antibodies may gain specificity (26,27). In these antibodies, a set of
complementarity determining region (CDR) residues were found to interact with the phosphate
group, while others make contacts with the amino acid residues nearby, leading to recognition of
both the modification and target sequence. Therefore, balancing the stability of interaction of
these distinct sets of CDR residues seems to be important in gaining both the affinity and
specificity. It is plausible that high-affinity PTM-targeting antibodies can achieve such tight
binding by stabilizing the interaction towards either the modified amino acid or peptide sequence,
potentially leading to unwanted cross-reactivity. Therefore, we hypothesized that a trade-off
between affinity and specificity might exist in PTM-targeting antibodies.
To test this hypothesis, we sought to assess the impact of in vitro affinity maturation on the
specificity of PTM-targeting antibodies. Starting from an existing high-specificity antibody (27)
targeting phosphorylated T231 of human tau (pT231, numbering based on the 441 amino acid
2N4R isoform), we generated a single-chain variable fragment (scFv) and improved its affinity
using directed evolution. To assess the specificity of PTM-targeting antibodies, we developed an
experimental approach based on yeast surface display (35) to quantify scFv binding against a
defined set of structurally similar epitopes. Specifically, peptides representing the
phosphorylated epitope, non-phosphorylated epitope, or a phosphorylated epitope with the same
amino acid composition, but scrambled sequence were labeled for detection. Their interactions
29
with yeast displayed antibody fragments were quantified using flow cytometry. Indeed, the
original high-specificity phospho-tau scFv showed nearly perfect specificity in this assay, in
agreement with previous characterizations (27). We found that through directed evolution, the
affinity could be improved by > 20-fold, reaching picomolar monovalent dissociation constants.
However, the majority of affinity-improved scFvs showed significant non-specific binding to the
non-phosphorylated epitope but not to the scrambled phospho-epitope, suggesting that high-
affinity clones tend to gain affinity by stabilizing the interaction with the amino acid residues
surrounding the modification site. Even though the majority of clones showed a trade-off
between their affinity and specificity, this was not necessarily the case for some unique clones.
By assessing the binding specificity, we identified a picomolar-affinity clone with specificity
profiles that match the original high-specificity antibody. These results demonstrated that a
quantitative assessment of both affinity and specificity was critical for identifying high affinity
and specificity antibodies targeting PTMs.
Materials and Methods
3.2.1 Yeast surface display of pT231 scFv
Amino acid sequence of VL and VH from an anti-tau pT231 antibody (27) was back-translated
to DNA sequence, yeast-codon optimized and synthesized (Integrated DNA Technologies). A
(G4S)3 linker was inserted between the VL and VH sequences (5’-VL-linker-VH) to create a
fusion construct encoding a scFv sequence. The synthesized scFv gene was cloned using PCR to
add a 5’ NheI and a 3’ BsrGI (New England Biolabs) restriction sites. The PCR products were
ligated into the vector pCT4RE (35) (a gift from Dr. Eric Shusta). This plasmid included an N-
30
terminal c-myc epitope tag and a C-terminal FLAG epitope tag. The pCT4RE plasmid was
transformed into S. cerevisiae EBY100 (35) (a gift from Dr. Dane Wittrup).
3.2.2 Peptide modification and purification
All peptides used were purchased from a commercial source (Peptide 2.0). Peptides contained a
C-terminal cysteine for chemical modification. Peptides to be modified were dissolved at 1 mM
in phosphate-buffered saline (PBS, 8 g/L NaCl, 0.2 g/L KCl, 1.44 g/L Na2HPO4, 0.24 g/L
KH2PO4, pH 7.4) at room temperature. A 10 mM stock solution of Alexa Fluor 488 C5
maleimide (Thermo Fisher Scientific) was prepared in deionized H2O, and a 100 mM stock
solution EZ-link maleimide-PEG11-biotin (Thermo Fisher Scientific) was prepared in dimethyl
sulfoxide (DMSO). The Alexa Fluor 488 C5 maleimide stock solution was protected from light
by wrapping container with aluminum foil. For peptide modification, either Alexa Fluor 488 C5
maleimide or EZ-Link maleimide-PEG11-biotin stock solution was added to the peptide PBS
solution at a final reagent concentration of 2 mM. The reactions proceeded at 4 oC overnight in
the dark.
Modified peptides were purified using an HPLC system (Shimadzu SCL-10Avp), equipped with
a preparative column (Waters, Atlantis T3, 100 Å, 10 µm, 10 mm x 250 mm). For the mobile
phase, solvent A consisted of 99.9 % H2O and 0.1 % trifluoroacetic acid, and solvent B consisted
of 99.9 % methanol and 0.1 % trifluoroacetic acid. Solvent flow rate was 3.5 mL/min. The
column temperature was set at 50 oC. For each HPLC run, 100 µL of the peptide reaction mixture
was injected. The gradient for peptides was as following: 0 % solvent B at 0 min, 20 % solvent B
at 1 min, 40 % solvent B at 26 min, solvent 75 % B at 31 min, and 100 % solvent B at 36 min.
Purified peptide fractions, which were detected at 214 nm, were collected, lyophilized, and
stored at - 20 oC.
31
Mass spectrometry of the modified peptides was conducted using a QStar Elite hybrid QTOF
mass spectrometer (AB Sciex). TOF spectra ((300-2000 m/z) were acquired using Analyst QS
2.0 software.
3.2.3 Directed evolution of pT231 scFv
pT231 scFv was randomly mutagenized as described previously (36) with the following
modifications. Error-prone PCR using a mixture of triphosphates of nucleoside analogs. Each
PCR consisted of ~ 250 ng of plasmid template, 0.5 µM of forward and reverse primer, 200 uM
of dNTP mix (Thermo Fisher Scientific), Taq PCR buffer (without MgCl2), 2 mM of MgCl2, 2.5
units of Platinum Taq DNA polymerase (Thermo Fisher Scientific), 2 µM of 8-Oxo-2'-
deoxyguanosine-5'-triphosphate (Tri-Link Biotechnologies, 8-Oxo-dGTP) and 2 µM of 2'-
Deoxy-P-nucleoside-5'-Triphosphate (Tri-Link, Biotechnologies dPTP) in a 50 µL of reaction
volume. The mixture was denatured at 94 oC for 3 min followed by 19 cycles of 94 oC for 45 s,
55 oC for 30 s, and 72 oC for 60 s and a final extension of 72 oC for 10 min on a thermocycler.
The PCR product was gel-purified using a gel extraction kit (Zymo Research). 2 µL of purified
mutagenesis product was added to a 100 µL of PCR reaction solution without using nucleoside
analogs to amplify the mutated fragments. About 6.4 µg of mutated pT231 scFv gene, and 1 µg
of vector pCT4RE (a gift from Dr. Eric Shusta) were combined with 200 µL of electrocompetent
EBY 100 (a gift from Dr. Dane Wittrup) and electroporated at 2.5 kV and 25 µF capacitance
using the GenePulser (Bio-Rad, 0.2 cm Electrode gap cuvette). The electroporated cells were
grown in SD-CAA media (0.1 M sodium phosphate, pH 6.0, 6.7 g/L yeast nitrogen base, 5 g/L
casamino acids, 20 g/L glucose) at 30 oC and 250 rpm. Frozen aliquots of the library were
prepared by freezing 10x library diversity in yeast freezing media (2 % glycerol and 6.7 g/L
yeast nitrogen base) at - 80 oC for long-term storage.
32
For the following rounds of gene diversification, it was realized by shuffling the library screened
in the previous round, followed by the above random mutagenesis procedure to incorporate
additional mutations. The plasmid DNA from the library pool was isolated using a yeast plasmid
miniprep kit (Zymo Research). pT231 scFv library gene was PCR amplified with primers
flanking the region. 2-4 µg of the PCR product was digested with 0.0015 unit of DNase I (New
England BioLabs) per µL in 100 µL of reaction mixture for 15 min at room temperature.
Fragments of 10-50 bp were purified from 2 % low melting agarose gel by electrophoresis and
then re-suspended in 50 µL of PCR reaction without primers. A PCR program of 94 oC for 3 min;
29 cycles of 94 oC for 30 s, 55 oC for 30 s, 72 oC for 30 s, and 72 oC for 10 min was used on a
thermocycler. Mixtures of DNA fragments were reassembled into a gene at defined positions
based on gene homology.
Fluorescence activated cell sorting was used to screen for yeast cells displaying high-affinity
scFvs. 10x library diversity of cells were pelleted and washed with 1 mL of phosphate-buffered
saline with bovine serum albumin (PBSA, 8 g/L NaCl, 0.2 g/L KCl, 1.44 g/L Na2HPO4, 0.24 g/L
KH2PO4, 1 g/L bovine serum albumin, pH 7.4) twice. The cells were incubated with Pphos-b in 3
mL PBSA at room temperature for 1 hour. After labeling with peptide, the cells were primarily
stained with mouse anti-FLAG (Sigma, 1:1000 dilution) in 1 mL of PBSA on ice for 30 min, and
then secondarily stained with goat anti-mouse Alexa Fluor 488 (Thermo Fisher Scientific, 1:100
dilution) and streptavidin R-phycoerythrin (SAPE, Thermo Fisher Scientific, 1:100 dilution) in 1
mL of PBSA on ice for 30 min. Pelleting and washing the cells with 500 µL of PBSA were
required between all labeling steps to remove any unbound peptides and antibodies. After
labeling, the cells were re-suspended in 3 mL of PBSA and treated with 1 µM of unmodified
phosphopeptide at room temperature for 7 min. The screening was carried out on a Becton-
33
Dickinson FACSAria II, which was equipped with five different lasers. Data of cell screening
was acquired using BD FACSDiva software. The collected cells were grown in SD-CAA media
plus 1 % kanamycin at 30 oC, 250 rpm overnight. After several rounds of cell propagation, 10x
library diversity of cells were archived in yeast freezing media at -80 oC.
3.2.4 Purification of pT231 scFvs
The pT231 scFv gene was inserted into the vector pRS316 (a gift from Dr. Eric Shusta), which
had been restriction-digested with NheI and BsrGI (New England Biolabs). This plasmid
included a C-terminal hexa-histidine epitope tag. The pRS316 plasmid was transformed into S.
cerevisiae YVH10 cells (37) (a gift from Dr. Eric Shusta) for protein secretion and purification.
Cells were grown in 500 mL of SD-CAA plus tryptophan media at 30 oC, 250 rpm for 3 days,
pelleted, and re-suspended in 500 mL of SG-CAA plus tryptophan with 0.1% bovine serum
albumin (Sigma-Aldrich) at 20 oC, 250 rpm for 3 days to induce and secrete the scFvs. The cells
were pelleted, and supernatant containing scFv was collected and filtered using bottle-top filter
(Corning, PES membrane, 0.22 µm) to remove the yeast cells and cell lysate. The scFv
supernatant was firstly concentrated to 50 mL by ultrafiltration using a selective permeable
membrane (Millipore Sigma, 10 kDa) under nitrogen pressure. The concentrated supernatant was
then dialyzed to exchange buffer in 5 L of phosphate-buffered saline (0.01 M sodium phosphate,
0.15 M NaCl, pH 8.0) at 4 oC overnight. The dialyzed scFvs solution was purified as described
previously (38), using the SuperFlow Ni-NTA beads (Qiagen). The purified scFvs were resolved
using SDS-PAGE in tris-glycine running buffer (14.4 g/L glycine, 1 g/L SDS, 3.03 g/L Tris base)
with 12 % Bis-Tris gels. SeeBlue® pre-stained protein standard (Thermo Fisher Scientific) was
used to determine the molecular weights.
34
3.2.5 Fluorescent tagging of purified pT231 scFvs
The scFv elution fractions were pooled and concentrated using a centrifugal filter unit (Millipore
Sigma, Amicon Ultra, 10 kDa MWCO). The concentrated scFvs were diluted to 3 mg/mL in 0.1
M sodium bicarbonate buffer at pH 8.3. The amine-reactive dye Alexa Fluor 594 NHS ester
(Thermo Fisher Scientific) was prepared in dimethylformamide (DMF) at 10 mg/mL
immediately before use. 35 µg of Alexa Fluor 594 NHS ester from the stock solution was added
to 100 µg of purified scFv. The reaction mixture was incubated for 1 hour at room temperature in
the dark. Unreacted dye was removed using the Zeba desalting columns (Thermo Fisher
Scientific, 7 kDa MWCO), and the concentrations of modified scFvs were determined using a
Bradford assay (Thermo Fisher Scientific).
3.2.6 Immunocytochemistry in HEK 293 cells
The HEK 293 cells were transfected with pRK5-EGFP-Tau (Addgene 46904), which contained
human tau (MAPT 0N4R) fused to EGFP at the N-terminus. Cells were rinsed once with 0.5 mL
of ice-cold PBSCM buffer (0.15 M NaCl, 1.9 mM NaH2PO4, 8.1 mM Na2HPO4, 1 mM CaCl2,
0.5 mM Mg2SO4, pH 7.4), fixed in 4% paraformaldehyde (Sigma) in PBSCM at room
temperature for 15 min, and then permeabilized using 0.1% Triton X-100 (Acros Organics) in
PBSCM at room temperature for 15 min. After washing three times with PBSCM, the cells
incubated with 0.1 µM of Alexa Fluor 594 conjugated scFv in 0.5 mL of PBSCM with 40% goat
serum (PBSG) at 4 oC overnight in the dark. The cells were washed three times with ice cold 0.5
mL PBSCM, stained with 0.3 µM of DAPI (Thermo Fisher Scientific) at room temperature for 5
min, washed once with 0.5 mL of ice-cold PBSCM. The cells were imaged using a Nikon A1R
confocal microscope (UConn Center for Open Research Resources and Equipment).
35
3.2.7 Immunohistochemistry in brain tissue sections
All mouse (n=3 PS19, 3 wild-type C57BL6/J, and 3 tau knock-out) and human (n=2 AD cases
and n=2 aged control) tissues were sectioned at 30 µm stained as free floating sections using
Netwell baskets (VWR Cat#29442-132) in 12-well Falcon plates (Cat#353043). Extracted mouse
tissues were flash frozen on dry ice, sectioned onto slides (Millenia 2.0 Adhesion Slides Catalog
# 318), and fixed in cold 4% PFA for 3 minutes prior to staining. Human tissues were fixed and
stored in PLP until sectioned using a Leica VT1200S vibratome. To quench lipofuscin
autofluorescence, human tissue sections were photobleached under a 1500 lumen white LED
bulb for 72 hours at 4°C while suspended in PBS with no lid or covering (39). Following mouse
tissue fixation and human tissue bleaching, all sections were then washed three times in PBS for
30 seconds per wash followed by a 5-minute incubation in detergent media (TBS with 0.25%
Triton-X). Tissue was then incubated for 30 minutes in citrate-based antigen unmasking solution
(Vector Cat#H-3300) at 95°C. Tissue was blocked with gentle rotating for 2 hours in detergent
media with 5% donkey serum (Sigma-Aldrich D9663-10mL). After blocking, free floating
human tissue sections were moved into separate wells of a 24-well plate with 300 µL of the
appropriate scFv dilution and no basket; mouse tissue was stained directly on the slides with 200
µL of the appropriate scFv dilution. Tissues were incubated with the scFvs for 3 hours at room
temperature with gentle rotation.
Following scFv incubation, tissue sections were incubated with 1 µg/mL of DAPI in the dark for
5 minutes at room temperature. Tissues were then washed four times in PBS for 5 minutes each
and carefully placed on Millenia 2.0 slides using a histology brush if free floating (Fisher
Scientific Cat#NC0344756). Excess water was allowed to evaporate for 15 minutes, then each
slide was mounted with 100uL of Prolong Gold Antifade Reagent (Thermo Fisher Cat#P36930)
36
and a 1 mm cover slip. Imaging was done using a Zeiss LSM 710 confocal microscope, ensuring
equal parameters for digital gain (790), pinhole (30 um), and laser power (4.4%) for all images.
Following imaging, images were processed using the Fiji distribution of ImageJ. The magic
wand tool was used to select individual tangles as a region of interest (ROI) which was then
measured for average fluorescence intensity
Results
3.3.1 Yeast surface display of a phospho-tau scFv
To test our hypothesis, we first developed an assay to quantify antibody specificity using yeast
surface display (Figure 3.1). Synthetic peptides that represent the exact target epitope, as well as
other potentially cross-reacting species were labeled for detection. Since yeast surface display
allows surface expression of 40,000-50,000 scFvs per cell (35,40), we anticipated that binding of
multiple structurally-related epitopes displayed on yeast could be quantified using multicolor
flow cytometry (Figure 3.1). To demonstrate this concept, we used the previously identified
high-specificity anti-phospho-tau antibody which targets pT231 (27). We cloned the antibody as
a VL-VH single-chain variable region fragment (pT231 scFv), fused to the N-terminus of the
yeast cell wall protein Aga2p (Figure 3.1).
37
Figure 3.1 Schematic representation of antibody specificity quantification using yeast surface display Individual peptide epitope binding to single chain variable region fragments displayed on yeast can be quantified. By labeling multiple peptides with distinct tags, competitive binding of potentially cross-reacting epitopes, such as a phosphopeptide and non-phosphopeptide (peptide mix indicated by a dotted box) can be quantified using multicolor flow cytometry.
3.3.2 Chemical labeling of peptides to generate distinctively labeled epitopes
To detect the interaction between pT231 scFv and the target phosphoepitope, a synthetic
phosphopeptide (Pphos, KKVAVVRpTPPKpSPSSAKC) was labeled with Alexa 488 at the C-
terminal cysteine using Alexa Fluor 488 C5 maleimide. The resulting peptide was purified using
high-performance liquid chromatography (HPLC), and the label was validated using mass
spectrometry (Figure 3.2a). To quantify the specificity of the scFv to structurally similar
epitopes, including the non-phosphoepitope and a phosphoepitope with scrambled amino acid
sequence, we labeled peptides representing the non-phosphoepitope (Pnonphos,
KKVAVVRTPPKSPSSAKC) and a scrambled phosphoepitope (Pscram,
KAVpSASVPVSRKpTKPPKC) with biotin at the C-terminal cysteine using maleimide-PEG11-
biotin (Figure 3.2, data not shown). To enable measurement of binding curve, we also generated
a phosphopeptide labeled with a second tag, biotin (Figure 3.2b).
38
Figure 3.2. Modified peptides purification and confirmation a, conjugation of Alexa Fluor 488 C5 maleimide to cysteine-Pphos and HPLC purification. HPLC elution profiles of free cysteine-Pphos (top), free Alexa Fluor 488 C5 maleimide (second), and conjugation reaction of the cysteine-Pphos to Alexa Fluor 488 C5 maleimide (bottom). A good separation between the Pphos-A488conjugate and other reactants facilitates purification. b, mass spectra showing Pphos-b and Pnonphos-b conjugate with no free cysteine-peptide.
3.3.3 Verification of binding activity of yeast displayed pT231 scFv
The scFv expression on the yeast cell surface and binding to the phosphoepitope were confirmed
using confocal microscopy (Figure 3.3a), using Streptavidin-R-phycoerythrin (SA-PE) as a
detection reagent. The affinity (dissociation constant Kd) of the yeast displayed pT231 scFv, as
determined by titrating the biotinylated phosphopeptide (Pphos-b) was 2.2 ± 0.4 nM (Figure
3.3b, mean ± standard deviation), which is lower than the monovalent affinity for the full-length
IgG (Kd = 0.35 nM) generated using the same variable regions (27). Even though scFv constructs
are highly advantageous for imaging (41) and delivery (42-44), a 5 to 15-fold reduction in
39
affinity upon conversion of IgG to scFv had been reported (42,45), potentially due to higher
stability and functional concentration of full IgGs.
Figure 3.3 Binding activity of yeast displayed pT231 scFv a, confocal microscope images of yeast displayed pT231 scFv and an isotype-matched control scFv. scFv expression shown in red was detected using a mouse anti-FLAG antibody to assess full-length scFv expression. Cognate phosphopeptide (Pphos) binding is indicated in green. Scale bars, 5 µm. b, affinity measurement of yeast displayed pT231 scFv. The yeast cells were incubated with serial dilution of Pphos. Error bars indicate standard deviation from three independent experiments.
To confirm that the yeast displayed scFv was properly folded, we generated alanine point
mutations in the CDR residues that interacted with the phosphoepitope (Figure 3.4), according
to the published x-ray structure of the antibody-phosphopeptide complex (27). Alanine point
mutations in all phosphoepitope interacting CDRs (single alanine substitutions in either CDR-H2,
CDR-H3, CDR-L1, CDR-L2, or CDR-L3) substantially reduced phosphopeptide binding
(Figure 3.4), indicating that CDR residues were correctly presented in the yeast displaying scFv.
40
Figure 3.4 Assessment of CDR mutations on phosphopeptide binding of yeast displayed pT231 scFv Alanine point mutations were introduced to scFv residues that directly interact with the phosphopeptide, according to the pT231 Fab-phosphopeptide crystal structure (PDB entry 4GLR). Yeast displayed wild-type pT231 scFv and alanine point mutants at indicated amino acid positions were incubated with Pphos-b at 1 µM. Control scFv indicates an isotype-matched random scFv. Amino acid numbering and CDRs are defined according to Kabat numbering scheme. Error bars indicate standard deviation from three independent experiments. Dunnett's multiple comparisons test, * P ≤ 0.05, **** P ≤ 0.0001, otherwise, P > 0.05.
3.3.4 Assessment of antibody phospho-specificity using yeast surface display
Having verified the binding activity of the pT231 scFv, we quantified the specificity of the scFv
to structurally similar epitopes, including the non-phosphoepitope and a phosphoepitope with
scrambled amino acid sequence. When we assessed the binding of pT231 to the phosphopeptide
labeled with Alexa 488 (Pphos-A488) using flow cytometry, we found that the signal intensity of
scFv-phosphopeptide was lower, compared to the signal obtained from Pphos-b (data not shown).
Such a difference could be attributed to the fact that the brightness of R-phycoerythrin (molar
absorption coefficient ´ quantum yield = 19.6 ´ 105 M-1cm-1 ´ 0.82 = 16.1 ´ 105 M-1cm-1) (46) is
41
24-fold higher than that of Alexa 488 (0.73 ´ 105 M-1cm-1 ´ 0.92 = 0.67 ´ 105 M-1cm-1,
manufacturer specifications). Therefore, we amplified the signal from Alexa 488, using an anti-
Alexa 488 antibody (rabbit polyclonal) followed by an anti-rabbit IgG conjugated with Alexa
488, which resulted in 10-fold increase in the scFv-Pphos-A488 signal (data not shown). When
yeast cells displaying the pT231 scFv were incubated with an equimolar mixture of Pphos-b and
Pphos-A488 (both at 0.5 µM), signals from both labels were detected using flow cytometry
(Figure 3.5), as anticipated in the multi-color labeling scheme (Figure 3.1). To assess the
specificity, yeast cells displaying pT231 scFv were incubated with a mixture of either Pphos-
A488 and Pnonphos-b (both at 0.5 µM) or Pphos-A488 and Pscram-b (both at 0.5 µM). In these
experiments, the yeast cells showed labeling almost exclusively by the target phosphopeptide
(Pphos-A488), and no detectable binding to the non-phosphopeptide (Pnonphos-b) or scrambled
phosphopeptide (Pscram-b) (Figure 3.5). These results confirmed the previous report of high
specificity of the pT231 antibody (27), and demonstrated the effectiveness of our specificity
quantification assay.
Figure 3.5 Assessment of the specificity of yeast displayed pT231 scFv Flow cytometry dot plots for yeast displayed WT pT231 scFv (top row) or an isotype-matched control scFv
42
(bottom row). The first column shows data for scFv expression and biotinylated phosphopeptide (Pphos-b) binding. In the following columns, yeast cells were incubated with a mixture of either Pnonphos-b and Pphos-A488 (second column), or Pscram-b and Pphos-A488, or Pphos-b and Pphos-A488, all at 0.5 µM.
3.3.5 Directed evolution of pT231 scFv for improved affinity
We performed directed evolution of the pT231 scFv by applying selection pressure to enhance
the affinity (Figure 3.6a). We generated a random mutation library of the pT231 scFv using
error-prone PCR (47) followed by electroporation in yeast (48), yielding approximately 3 ´ 107
transformants displaying the scFv variants. The scFv library was subjected to three rounds of
fluorescence-activated cell sorting (FACS) (Figure 3.6a). After round 1 and 2, the sorted scFv
sequences were further diversified using DNA shuffling followed by random mutagenesis
(Figure 3.6a). Throughout the FACS process, selection pressure for higher affinity was applied
by incubating yeast cells displaying pT231 mutants with progressively lower concentration of
Pphos-b (Figure 3.6a) followed by incubation with 1 µM of unlabeled phosphopeptide (Pphos)
for dissociation (49), and sorting for high Pphos-b binding vs. scFv expression (using anti-c-Myc
antibody) (Figure 3.6b). After three rounds of FACS, a population of high Pphos-b was enriched
(Figure 3.6c).
43
Figure 3.6 Directed evolution of yeast displayed pT231 scFv variants with improved affinity a, flow chart for the directed evolution process. High-affinity variants were sorted using FACS. b, flow cytometry dot plot with a sorting gate used for screening cells with scFv expression and high Pphos-b binding. c, Dot plots comparing yeast displaying WT pT231 scFv (left) and a yeast pool resulting from three rounds of sorting and mutagenesis (right) were labeled for scFv expression and Pphos-A488 binding.
We sequenced individual clones from this pool and identified 27 unique scFv mutant sequences
(Figure 3.7). Even though all 27 clones had distinct sequences with mutations throughout the
scFv sequence, several high occurrence mutations were found (Figure 3.7).
44
Figure 3.7 Amino acid sequence of individual mutants isolated from the screen for improved affinity The sequences in the boxes indicate CDRs. High occurrence mutations are indicated in bold. Kabat numbering is indicated at the top.
The majority of frequent mutations (residues with occurrence labeled in bold in Figure 3.7) were
found in or near the CDRs (Kabat numbering): VLW50L and VLS56C (L2), VLA95T,
VHG100HD (or S), VHD101G, and VHA102T (H3) (Figure 3.8). Mutations VLD57G, VHG74C,
and VHA93V were in the framework regions (Figure 3.8).
45
Figure 3.8 High occurrence mutations identified from the screen mapped on WT pT231 scFv structure Heavy chain is shown in green and light chain in purple. Position of high occurrence mutations are indicated in red, and labeled with the wild-type amino acid. Boxed amino acids indicate those mutated in the scFv 3.24. The structure was generated using CCP4MG, based on protein data bank (PDB) entry 4GLR.
3.3.6 Characterization of affinity and specificity of scFv mutants
To assess the effectiveness of the directed evolution, we characterized the clones isolated after
three rounds of sorting for their affinity and specificity. For these analyses, we chose the first 14
unique clones identified from sequencing (clones 3.1 to 3.14, Figure 3.7). To rapidly compare
the affinity to that of the WT pT231 scFv, we incubated yeast cells displaying mutant scFvs with
2 nM of Pphos-b, which was near the Kd of the WT pT231 scFv. At this peptide concentration, 7
out of 14 clones showed significantly higher binding (normalized to the scFv expression level)
46
compared to that of the WT scFv (Figure 3.9a, P < 0.05, Dunnett’s multiple comparisons test).
Based on the binding/expression level, we chose scFv mutant 3.05 (containing 6 mutations,
Figure 3.7) and measured its affinity by displaying it on yeast and generating a titration curve
for the Pphos-b binding (Figure 3.9b). scFv 3.05 showed a 21-fold improvement in affinity over
that of the WT pT231 scFv, reaching picomolar monovalent binding affinity (Kd = 92 ± 1 pM)
(Figure 3.9b). To characterize the phospho-specificity of the 14 mutant clones, we first
measured the individual binding to Pnonphos-b and Pscram-b of the 14 clones (Figure 3.9c, d).
We found that of the 14 clones, 6 showed significant binding to the Pnonphos-b (Figure 3.9c, P
< 0.01, Dunnett’s multiple comparisons test), while none of the clones showed detectable
binding to the Pscram-b (Figure 3.9d). When we assessed the specificity of the high-affinity
scFv 3.05 using competitive binding between Pphos-A488 and Pnonphos-b, significant binding
to Pnonphos-b was detected (data not shown). Taken together, these results suggested that some
of the clones gained affinity by stabilizing the interaction to the target peptide sequence, while
none of the clones gained affinity to the phosphate group alone to cause cross-reactivity towards
Pscram-b. Conversely, the fact that some of the clones did not show significant binding to the
Pnonphos-b (Figure 3.9c) suggested that improving affinity did not necessarily lead to enhanced
cross-reactivity towards the non-phosphorylated epitope. In order to identify an optimal clone
with improved affinity and high specificity, we selected additional clones based on the sequence
homology and occurrence of mutations (Figure 3.7), and assessed their binding to Pphos-b,
Pnonphos-b, and Pscram-b. From this characterization, we identified scFv 3.24 (containing 5
mutations) that showed 11-fold of improvement in binding affinity over the WT pT231 (Figure
3.9e, Kd = 160 ± 50 pM) and no significant change in binding to Pnonphos-b compared to that of
wild-type pT231 (Figure 3.9f). These results indicate that in phospho-specific antibodies,
47
applying selection pressure towards improved affinity does result in cross-reactivity in a
significant fraction of clones. Therefore, high-affinity clones need to be checked for phospho-
specificity in order to improve the binding affinity through directed evolution.
Figure 3.9 Characterization of affinity and specificity of individual scFvs a, binding intensity normalized to the expression level of yeast displayed WT pT231 scFv and mutants to Pphos-b at 2 nM. The right most sample was the WT pT231 scFv incubated with Pphos-b at 1 µM, as a positive control. b, Affinity titration for yeast-displayed scFv 3.05. Binding intensity of yeast displayed WT
48
pT231 scFv and mutants to Pnonphos-b at 1 µM in c, and Pscram-b at 1 µM in d, respectively. e, affinity titration for yeast-displayed scFv 3.24. f, Binding intensity normalized to the expression level of yeast displayed WT pT231 scFv and mutants to Pnonphos-b at 1 µM. Error bars indicate standard deviation from three independent experiments. Statistics for panels (a, c, d, and f): Dunnett's multiple comparisons test, *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001, ****P ≤ 0.0001, otherwise, P > 0.05.
3.3.7 Cell and tissue section labeling using purified scFvs
High affinity and specificity reagents are expected to enable sensitive detection of tau
phosphorylation in complex samples. We tested the performance of the scFvs in
immunocytochemistry and immunohistochemistry experiments. In order to perform the labeling
experiments, we cloned the wild-type scFv and mutant 3.24 into a vector containing a synthetic
secretion signal (38) to secrete them using yeast S. cerevisiae. The secreted scFvs were purified
using a hexa-histidine motif at the C-terminus. When we resolved the purified scFvs using SDS-
PAGE followed by Coomassie staining, the gel loaded with purified scFvs showed a single
protein band with a molecular weight of 28 kDa, which is typical for scFvs (50), with ~92%
purity as quantified by densitometry of the gel (data not shown). To enable direct imaging of the
scFvs without the use of secondary reagents, the purified scFvs were fluorescently labeled with
Alexa 594 through modification of lysine residues (NHS-Alexa Fluor 594) and purified. In order
to assess the specificity, the fluorescent scFvs were used to label human embryonic kidney cells
(HEK293FT) transiently expressing human tau (microtubule-associated protein tau (MAPT)
0N4R) as a GFP fusion (EGFP-tau) (51). When high concentrations (100 nM) of the WT pT231
scFv or scFv 3.24 were used for staining, clear scFv signal was detected in HEK293FT cells
expressing wild-type tau (Figure 3.10, top row) but not to the same cells treated with 𝜆
phosphatase (Figure 3.10, second row). These results indicate that the binding of wild-type
pT231 scFv and mutant 3.24 requires phosphorylation of tau, and as reported previously, the tau
expressed in HEK293 cells is phosphorylated (52,53). In order to further assess the specificity of
49
the scFvs, we generated point mutants of human tau at the target phosphorylation site (T231)
either incapable of phosphorylation (T231A) or pseudophosphorylated (T231E). In HEK293FT
cells expressing EGFP-tau T231A, both the wild-type pT231 scFv and mutant 3.24 showed no
detectable binding (Figure 3.10, third row), while in the same cells expressing EGFP-tau T231E,
both scFvs showed clear binding (Figure 3.10, bottom row). These results indicate that the scFv
binding requires tau phosphorylation at T231.
Figure 3.10 Confocal microscopy images of HEK293FT cell labeled using purified pT231 scFvs Purified fluorescently labeled WT pT231 scFv and mutant 3.24 binding to HEK293FT cells expressing WT tau (top row), WT tau treated with lambda phosphatase (second row), tau T231A (third row), and tau T231E (bottom row). All human tau constructs were fused to EGFP at the N-terminus (EGFP-tau). The first column shows DAPI staining in blue. The second column shows tau expression in green. The third column shows scFv binding in red. Scale bars, 50 µm.
When we compared the labeling of WT and scFv 3.24 at lower concentrations, scFv 3.24 showed
significantly higher binding signal above background compared to that of WT at scFv
concentrations ranging from 80 pM to 50 nM (Figure 3.11, P < 0.05, student’s t-test).
50
Figure 3.11 Quantification of cell labeling fluorescence intensity using purified pT231 scFvs Fluorescence intensity of WT pT231 scFv and mutant 3.24 binding to HEK293FT cells expressing WT tau was measured at scFv concentrations from 16 pM to 50 nM. Error bars indicate standard deviation from three independent experiments. Statistics: Student’s t-test, *P ≤ 0.05.
We proceeded to test the fluorescently labeled scFvs in the staining of tissue sections from both
PS19 mouse tissue slices and human AD tissue slices; PS19 mice expressed the aggregation-
prone mutant P301S of human tau (1N4R) under the mouse prion protein (Prnp) promoter (54).
In brain slices of 8-month old PS19 mice, wild-type pT231 scFv and mutant 3.24 demonstrated
staining patterns characteristic of neurofibrillary tangles (Figure 3.12a), suggesting specific
labeling of tau. This staining was dose-dependent for both scFvs (Figure 3.12b). Neither scFv
showed detectable staining in either tau knock-out or WT C57Bl/6 mice (Figure 3.12c). Staining
of human AD tissue also revealed characteristic tangle staining (Figure 3.12d) with no
detectable staining in aged control brain tissues. However, in the human tissues the mutant scFv
3.24 showed significantly higher signal than the WT scFv at the 50 nM working concentration
(Figure 3.12e), suggesting higher affinity and more robust staining for the mutant scFv. These
51
results further demonstrate that the high-affinity antibody generated in this study has
exceptionally high specificity for phospho-tau.
Figure 3.12 Tissue labeling using purified pT231 scFvs a, immunohistochemistry using the scFvs on brain slices of 8-month old mice expressing P301S mutant tau indicates that both scFvs robustly identify neurofibrillary tangles in tissue. Each scFv was used at 50 nM and 10 nM to identify optimal staining concentrations and show a dose response for staining intensity, quantified in b. tau is shown in red with nuclei stained using DAPI in blue. The 10nM concentrations showed significant reduction in staining intensity for both the WT and 3.24 scFv (P ≤ 0.0001 by one-way ANOVA with P ≤ 0.0001 between the 50 nM and 10 nM working concentrations of each scFv using Tukey’s multiple comparisons test). There were no statistical differences between the WT and 3.24 scFv at either concentration. c, neither scFv stained WT or tau knockout mouse tissue at the 50 nM working concentration, indicating no off target staining or background fluorescence. d, immunohistochemistry of human AD and aged control tissues also indicate that both scFvs stain phospho-tau tangles in disease (scFv concentration = 50 nM). However, in the human tissue the high affinity scFv 3.24 shows a significant increase in staining intensity compared to the WT scFv, which is quantified in e (P ≤ 0.0001 using a two-sample t-test assuming unequal variances).
52
Discussion
In this study, we applied directed evolution to enhance the affinity of a phospho-specific
antibody, and quantified the impact of in vitro affinity maturation on specificity. High-affinity
antibody fragments with dissociation constants reaching picomolar range were successfully
generated. Based on the crystal structure of the WT pT231 antibody bound to the
phosphopeptide (27), only one mutation occurred in a residue that directly contacts the
phosphoepitope (Figure 3.8, VLW50L). Five out of 9 high occurrence mutations were located
near the direct contact sites (VLS56C, VLA95T, VHG100HD (or S), VHD101G, and VHA102T).
The pattern of mutations occurring in amino acid positions near, but not at the direct contact site
has been frequently observed in high affinity antibody variants (55,56). Such mutations may
contribute to enhanced affinity through preorganization and rigidification of the paratope (57-59).
Mutations VHG100HD, VHD101G, and VHA102T are also located at the VH-VL interface,
suggesting that stability of interaction between the two domains may contribute to affinity
improvement. Interestingly, high occurrence mutations found in the optimal variant 3.24
(VLD57G, VHG74C, and VHA93V) were exclusively located in the framework region. This
suggests that this variant may have gained affinity through optimization of structural plasticity
(60,61). Further investigation on the impact of individual mutations on affinity and specificity
may provide new insights on designing PTM-specific antibodies.
High-affinity reagents would be highly advantageous in detecting PTMs from complex samples.
Our results show that the high affinity variant scFv generated significantly higher signal
compared to that of the wild-type scFv in detecting tau from human tissue, but not in mouse
tissue. The discrepancy in staining intensities may be explained by differences in the expression
level of human tau and level of phosphorylation. In the transgenic mice PS19, the expression of
53
human tau P301S is driven by the prion protein promoter, which is strongly induced in neuronal
cells (62), leading to 3-5 fold higher levels of tau expression than endogenous levels (54).
Moreover, in PS19 mice, the solubility of tau decreases by 3 months of age, and becomes
hyperphosphorylated (54,63). Therefore, the level of insoluble aggregates of phosphorylated tau
in the mouse tissue may be significantly higher than in human AD patient tissue. Additionally, in
the mouse tissue, the labeling intensity was significantly diminished at a scFv concentration of
10 nM, suggesting a possibility for mass transfer limitation of the labeling reagent. Very slow or
limited antibody penetration has been reported for fixed samples (64). This shows the need for
high antibody concentrations in labeling fixed tissue, and highlights the importance of antibody
specificity.
While the scFv identified in this study can be converted to full-length IgGs, the scFv construct
provides unique advantages for detecting tau due to its small size. Compared to full-length IgGs
(150 kD), scFvs are approximately one-sixth in molecular weight (25 kD). This has led to
significant enhancement in processes that rely on diffusion, such as tissue penetration. For
example, when the blood-brain barrier was transiently opened through focused ultrasound, scFvs
showed 5- to 57-fold higher uptake into the central nervous system compared to IgGs (44). In a
mouse model of tauopathy, intravenously injected scFvs were detected in the brain tissue in
significantly higher levels compared to that of IgGs (41). Moreover, scFv constructs enable
wider modes of delivery since they are encoded by a short single gene that can be packaged in
viruses. For example, intracerebroventricular injection of adeno-associated viruses carrying an
anti-tau scFv resulted in reduced tau accumulation in a mouse model of tauopathy (43). Finally,
we demonstrated cell and tissue staining using scFvs directly labeled with a fluorophore. These
54
reagents have the potential to enable sub-diffraction limit imaging of phospho-tau due to greatly
reduced linkage error, which limit image resolution (65).
Phosphorylation of tau at T231 is a widely recognized modification for its association with AD.
In vitro and in cultured cells, pT231 decreases the ability of tau to bind and stabilize
microtubules (66,67). pT231 seems to occur early in AD progression and precedes paired helical
filament (PHF) formation (68). pT231 was found in the cerebrospinal fluid (CSF) of patients
with mild cognitive impairments who eventually developed AD (69). In addition, levels of
pT231 in the CSF were correlated with rates of hippocampal atrophy in AD patients (70). An
immunoassay of pT231 in the CSF demonstrated a 85% sensitivity (detection in 23 out of 27 AD
samples) and a 97% specificity (detection in 1 out of 31 non-AD samples) in AD brain extracts
(71). The high affinity and specificity pT231 tau antibody developed in this study is expected to
enhance the sensitivity and enable earlier detection of pT231.
A major finding of this study is that in vitro affinity maturation of the phospho-specific antibody
led to an increase in unwanted cross-reactivity towards the non-phosphorylated target sequence
in a majority of high-affinity variants. Considering the fact that the area of interaction available
from a modified amino acid is limited, stabilization of interaction with the nearby amino acid
sequence may be a common phenomenon in high-affinity PTM-specific antibodies. In a recent
study, monoclonal antibodies generated in guinea pig against pT18 of p53 showed picomolar
affinity (Kd = 0.20 nM) but also significant binding to the non-phosphorylated p53 (Kd = 130 nM)
(72). If such antibody is used at concentration ranges commonly used for cell or tissue staining
(10 - 50 nM), false positive signal from cross reactivity towards the non-modified target may be
detected. This illustrates an important point that antibody specificity is a function of antibody
concentration. Therefore, existing PTM-specific antibodies should be validated for their
55
specificity under varying concentrations. We suggest setting an upper concentration limit for
each PTM-specific antibody to prevent false positive detection. Until now, efforts to validate and
enhance specificity has been rarely made. Non-specific antibodies would be a major factor in
irreproducibility in biological experiments. Ongoing efforts to standardize ways to quantify and
document specificity of binding reagents (73) need to consider these aspects of antibody
specificity.
We introduced an approach using yeast surface display to quantify the specificity of PTM-
targeting antibodies. This enabled us to find that even among antibody variants selected only for
improved affinity, a fraction of clones maintained high specificity. This specificity quantification
approach may be widely adopted due to the general applicability of yeast surface display and its
compatibility with flow cytometry. We demonstrated the use of this approach for a phospho-
specific antibody, but it is readily applicable to other PTM-specific antibodies. In principle, this
approach would be suitable for assessing the specificity of affinity reagents in general. The
identified high-affinity variant showed exceptional specificity, as validated by cell and tissue
staining experiments. Due to the compatibility with FACS, this approach has potential to enable
directed evolution of antibody specificity. In particular, it would be useful in engineering the
specificity of antibodies with known cross-reactivity. For example, a recent study showed that a
significant fraction of commercially available PTM-specific tau antibodies has cross reactivity
(34). Directed evolution may be applied to reduce the cross-reactivity of these antibodies.
Conclusion
In this study, we introduced an approach using yeast surface display to quantify the specificity of
phospho-specific antibodies. We demonstrated the use of this approach for a phospho-specific
56
antibody, but it is readily applicable to other PTM-specific antibodies. In principle, this approach
would be suitable for assessing the specificity of affinity reagents in general. We also developed
a screening assay based on it to generate PTM-specific antibodies with improved affinity and
high specificity. We demonstrated that improving PTM- specific antibody affinity through
directed evolution results in cross-reactivity to the non-modified target sequence, there’s a trade-
off between affinity and specificity in PTM antibodies. However, quantifying specificity leads to
identification of a variant with high affinity and specificity.
The results from this study demonstrate that in vitro affinity maturation is a viable option for
improving the affinity of PTM-specific antibodies. In this process, it is essential to quantify the
specificity, particularly to the non-modified target sequence. By quantifying the specificity, it is
possible to apply selection pressures for improved affinity and specificity simultaneously.
57
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Introduction
Site-specific phosphorylation of the microtubule-associated protein tau is a well-defined
molecular signature linked to normal tau function as well as pathology in neurodegeneration.
Antibodies binding to phosphorylated tau (phospho-tau) are extensively used in research and
being adopted in clinical diagnosis (1-6). According to antibody databases (Antibodypedia and
Alzforum), thousands of anti-tau antibodies are currently available, including >180 unique
phospho-tau antibodies. However, few validation data have been published (7-9), which
indicated an alarming degree of non-specific binding frequently observed in phospho-tau
antibodies. To prevent waste of resource and irreproducibility of results, a standard measure of
specificity, equivalent to dissociation constants (Kd) for measuring affinity, is urgently needed.
Ideally, this parameter should enable users to compare the specificity, identify the source of non-
specificity, and clearly understand what is measured.
Existing specificity characterization methods relied on immunoblotting approaches using either
synthetic peptides, cell lysates or extracted paired helical filament (PHF) tau (7-9). Synthetic
peptides allow binding measurement to exactly defined phosphorylation sites, but is prohibitively
costly to cover all phospho-sites across the full-length tau. Cell lysates allow detection of non-
specific binding to non-tau proteins (for example by using cell lysates from a tau knockout
mouse) (7), but cannot measure cross-reactivity to non-modified tau or other phosphorylation
sites. Extracted PHF-tau from AD brain homogenates allows characterizing antibodies that do
not recognize normal human adult tau and measuring the cross-reactivity to nonphospho-tau by
treatment with phosphatase, but cannot capture the phospho-site specificity (9). Steinhilb and
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colleagues have generated transgenic flies expressing tau with alanine point mutations at several
serine-proline and threonine-proline motifs of human tau (10). In this study, they used fly head
extracts from these transgenic lines to perform western blotting using phospho-tau antibodies. In
principle, this approach may be applied for validating phospho-tau antibodies. However, all of
these approaches are not ideal for quantitative measurement due to frequent signal saturation in
immunoblotting experiments (11). To fill this critical need, we introduce a parameter termed Φ
(12) that quantifies specificity of phospho-tau antibodies and establish a generalizable, robust
assay to measure it.
The approach leverages flow cytometry for quantitative, multi-dimensional analysis of small
changes in antibody binding to different cell populations. In a typical immunodetection
experiment, signal from specific and non-specific binding cannot be distinguished. We sought to
enable this distinction within a single sample, by generating two populations of cells with
distinct fluorescence reporters. In one population of cells, the target protein (i.e., tau) is
expressed as a fusion to the enhanced green fluorescent protein (EGFP). In another population,
tau containing a site-specific alanine mutation is expressed as a fusion to a near infra-red
fluorescent protein (iRFP). The two populations of cells are mixed, and labeled with a phospho-
specific antibody. For a phospho-site specific antibody, any binding to iRFP-positive cells is
non-specific binding, enabling the measurement of the fraction of non-specific signal among all
antibody binding signals. In order to ensure the measurement of cross-reactivity to non-
phosphorylated tau, the experiment is repeated by replacing alanine mutant expressing cells with
tau expressing cells (iRFP fused to wild-type tau) treated with the lambda protein phosphatase
(λPP). Analogous to the alanine mutation assay, binding to iRFP-positive cells is identified as
non-specific signal. Based on these measurements, the fraction of non-specific signal is
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calculated, which is subtracted from 1 to obtain Φ. Therefore, Φ indicates the fraction of specific
signal.
Using this measurement, we measured the specificity Φ of 7 commonly used phospho-tau
antibodies (AT270, AT8, AT100, AT180, PHF-6, TG-3, and PHF-1), in addition to others. Our
results show that 5 antibodies (AT8, AT180, PHF-6, TG-3, and PHF-1) have Φ values near 1,
indicating nearly perfect specificity. Antibodies AT270, AT100 showed weak but detectable
non-specific binding to non-tau proteins in cells, with lower Φ values. This resulted in
background fluorescence in flow cytometry and immunocytochemistry experiments. The non-
specific binding also resulted in false positive detection in western blots, evidenced by additional
bands appearing in cell lysates that do not contain human tau. To our knowledge, this is the first
quantitative measurement of antibody specificity that enables specificity comparison with
information on the source of non-specificity.
Materials and Methods
4.2.1 Cloning of recombinant constructs of tau and mutant tau
The following plasmids were obtained from Addgene: a mammalian expression vector pRK5
encoding EGFP tagged wild-type (13) human tau (pRK5-EGFP-tau, # 46904, RRID:
Addgene_46904), which contains four C-terminal repeat regions and lacks the N-terminal
sequences (0N4R), a plasmid encoding iRFP (piRFP670-N1, # 44457, RRID: Addgene_45457),
and a mammalian expression vector pcDNA3 encoding hemagglutinin tagged constitutively
active glycogen synthase kinase-3b (GSK-3β) (S9A) (pcDNA3-GSK3b, # 14754, RRID:
Addgene_14754). The EGFP in pRK5 vector was first replaced by iRFP to construct an iRFP
tagged WT tau by golden gate assembly (pRK5-iRFP-tau) (14). This assembly was performed
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using a Type IIs restriction enzyme BsaI (New England Biolabs # R3535) and T4 DNA ligase
(New England Biolabs # M0202) (see Table 4.1 for primers used). Seven alanine mutant
variants were generated in iRFP tagged tau to preclude phosphorylation. Alanine mutations were
introduced to either Thr181, Ser202, Thr212/Ser214 (double alanine mutant), Thr231, Ser262,
Ser396/Ser404 (double alanine mutant), or Ser404. For this, site-directed mutagenesis was
conducted using overlap extension PCR (15) using the Phusion DNA polymerase (New England
Biolabs # M0530) (see Table 4.2 for primers used). Residue numbering of tau is according to the
longest adult human brain tau isoform.
Table 4.1 Primers for Golden Gate assembly of plasmids
DNA template Primers pRK5-EGFP-Tau 5’- TCCGCTGAGCCCCG -3’
5’- CCATGGTGGCGACCATC -3’ piRFP670-N1 5’- ATGGTAGCAGGTCATGCC -3’
5’- CGGATCCGCTCTCAAGCGCGG -3’
Table 4.2 Primers used for site-directed mutagenesis
Mutant Primers T181A 5’- GAGCTGGGTGGTGCCTTTGGAGCGGGC -3’
5’- GCCCGCTCCAAAGGCACCACCCAGCTC -3’ S202A* 5’- AGTGCCTGGGGCGCCGGGGCTGC -3’
5’- GCAGCCCCGGCGCCCCAGGCACT -3’ T212A/S214A 5’- GTTGGAAGGGCCGGGGCGCGGGAGCGG -3’
5’- CCGCTCCCGCGCCCCGGCCCTTCCAAC -3’ T231A 5’- GACTTGGGTGGAGCACGGACCACTGCCACCTTCT -3’
5’- AGAAGGTGGCAGTGGTCCGTGCTCCACCCAAGTC -3’ S262A 5’- TTCAGGTTCTCAGTGGCGCCGATCTTGGACTTG -3’
5’- CAAGTCCAAGATCGGCGCCACTGAGAACCTGAA -3’ S396A/S404A§ 5’- CAGACACCACTGGCGCCTTGTACACGATCTC -3’
5’- GAGATCGTGTACAAGGCGCCAGTGGTGTCTG -3’ S404A 5’- AGATGCCGTGGAGCCGTGTCCCCAGAC -3’
5’- GTCTGGGGACACGGCTCCACGGCATCT -3’ Note: *Previous study has shown that AT8 recognition requires only Ser202 to be phosphorylated in tau (13). §S396A/S404A were obtained using the S404A mutant as a template to generate double mutations.
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4.2.2 Cell culture and transfection
The human embryonic kidney (HEK) 293FT cells a high transfection efficiency isolate of the
HEK293T cells were purchased from ThermoFisher (# R70007), and used without further
authentication. The cell line is not listed as a commonly misidentified cell line by the
International Cell Line Authentication Committee. HEK293FT cells were grown in DMEM
supplemented with 10 % fetal bovine serum. The cells were frozen at early passage and used for
less than 2 months in continuous culture. HEK293FT cells were seeded in 24-well plate in the
same number 2 days before plasmid transfection to reach 70 % confluency. The cells were then
transfected with plasmid using the Lipofectamine 3000 reagent (Invitrogen # L3000001)
according to the manufacturer’s instructions.
4.2.3 Tau antibodies
10 phospho-tau antibodies raised against 7 phosphoepitopes in tau were validated for their
specificity (Table 4.3). The following monoclonal antibodies were commercially available:
AT270 (pThr181), AT8 (pSer202/pThr205), AT100 (pSer212/pSer214), AT180 (pThr231),
PHF-6 (pThr231), 1H6L6 (pThr231), and anti-tau (pSer404). The following monoclonal
antibodies were a gift of Dr. Peter Davies: TG-3 (pThr231 and conformation-specific) and PHF-
1 (pSer396/pSer404). One purified rabbit polyclonal antibody anti-tau (pSer262) was purchased
from Abcam. Tau-5 (Tau210-241, Invitrogen # AHB0042, AB_2536235) is a monoclonal
antibody recognizing phosphorylated and non-phosphorylated tau isoforms. Purified single-chain
antibody variable fragment (scFv) 3.24 targeting pThr231 was reported previously (16).
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Table 4.3 Phospho-tau antibodies
Name Epitope Host/ Isotype
Source Dilution
Working conc.
𝚽𝐚𝐥𝐚 𝚽𝛌𝐏𝐏
AT270 pT181 Mouse/IgG1, k Invitrogen, # MN1050, RRID: AB_223651
1:100 2 µg/mL 0.80±0.06 0.99±0.01
AT8 pS202/pT205
Mouse/IgG1 Invitrogen, # MN1020, RRID: AB_223647
1:100 2 µg/mL 0.95±0.07 0.95±0.05
AT100 pT212/pS214
Mouse/IgG1, k Invitrogen, # MN1060, RRID: AB_223652
1:100 2 µg/mL N/A N/A
AT180 pT231 Mouse/IgG1, k Invitrogen, # MN1040, RRID: AB_223649
1:100 2 µg/mL 0.99±0.01 0.98±0.01
PHF6 pT231 Mouse/IgG1 Invitrogen, # 35-5200, RRID: AB_2533210
1:250 2 µg/mL 0.98±0.01 0.97±0.00
1H6L6 pT231 Rabbit/IgG Invitrogen, # 701056, RRID: AB_2532360
1:250 2 µg/mL 1.00±0.00 0.99±0.01
TG-3 pT231 Mouse/IgM Peter Davies RRID: AB_2716726
1:100 N/A 0.96±0.01 0.95±0.06
pS262 pS262 Rabbit/IgG Abcam, # ab131354, RRID: AB_11156689
1:500 2 µg/mL 0.32±0.02 0.37±0.01
PHF1 pS396/pS404
Mouse/IgG Peter Davies RRID: AB_2315150
1:100 N/A 0.98±0.00 0.98±0.03
pS404 pS404 Rabbit/IgG Abcam, # ab92676, RRID: AB_10561457
1:200 0.1 µg/mL 1.00±0.00 0.99±0.00
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4.2.4 Quantitation of antibody specificity
Here we define a generally applicable parameter to quantify antibody specificity, based on
general discussions regarding multi-partner protein interaction by Hall and Daugherty (1). If an
antibody Ab is capable of binding to n different partners X1, X2, … Xn as described by the
following reactions,
𝐴𝑏 + 𝑋012,4 𝐴𝑏𝑋0
𝐴𝑏 + 𝑋612,7 𝐴𝑏𝑋6
…
𝐴𝑏 + 𝑋812,9 𝐴𝑏𝑋8
the specificity of the interaction between Ab and Xi can be defined as Φ: where:
Φ: ≡𝐴𝑏𝑋:𝐴𝑏𝑋<8
<=0= 1 −
𝐴𝑏𝑋<¹:8<=0
𝐴𝑏𝑋<8<=0
From this definition, 0 < Φ: < 1, where 1 indicates perfect specificity (Ab only reacts with its
intended antigen Xi), and 0 indicates no interaction with Xi. We used the following formula to
quantify the specificity of phospho-tau antibodies:
ΦCDEFCDE =[𝐴𝑏 − 𝑇𝑎𝑢CDEF]𝐴𝑏 − 𝐴𝑔MEMNO
where 𝑇𝑎𝑢CDEF indicates phosphorylated epitope sequences of tau at a defined site, and 𝐴𝑔MEMNO
indicates overall potential partners including 𝑇𝑎𝑢CDEF . In our assay, we determined the
specificity using a three-color labeling scheme, where the mixture of HEK293FT cells
transiently expressing EGFP tagged WT human tau and iRFP tagged alanine mutant tau was
incubated with a fluorescence-labeled antibody. In this case, the overall specificity ΦNON can be
obtained by measuring the following parameter:
69
ΦP =[𝐴𝑏 − 𝐴𝑔NON]𝐴𝑏 − 𝐴𝑔MEMNO
where 𝐴𝑔NON indicates antigen with an alanine mutation at the target phosphorylation site.
Therefore, ΦP in this case indicates the fraction of total binding signal to the alanine mutant
(non-specific binding signal from cells expressing the alanine mutant). ΦNON is defined as:
ΦNON = 1 − ΦP
Therefore, ΦNON value of 1 indicates no cross-reactivity to the alanine mutant. Experimentally,
ΦP was determined by measuring the fluorescence intensity of antibody labeling from each cell
in flow cytometry experiments (detailed description on how Φ values were obtained is below).
For example,
ΦP =𝐹𝐿0𝐹𝐿6
where FL1 and FL2 are fluorescence intensity of antibody labeling from HEK293FT cells
transiently expressing alanine mutant tau and WT tau, respectively. Fluorescence intensity was
calculated by subtracting the background fluorescence measured on the same day from a control
antibody with matching isotype. Analogously, ΦSTTis defined by replacing 𝐴𝑔NON with lambda
protein phosphatase (λPP)-treated antigen (𝐴𝑔STT). ΦSTT value of 1 indicates no cross-reactivity
to the de-phosphorylated tau.
4.2.5 Measurement of 𝜱: validation of phospho-tau antibody specificity
To measure ΦNON of phospho-tau antibodies, the HEK293FT cells were co-transfected with
plasmids pRK5-EGFP-tau WT and pcDNA3-GSK3b using the lipofectamine method described
above. In separate wells, HEK293FT cells were co-transfected with plasmids pRK5-iRFP-tau
(alanine mutant) and pcDNA3-GSK3b. Cells were detached from 24-well plate using 0.05 %
trypsin-EDTA (Gibco # 25300-054) for 10 min at 37 oC, then pelleted and washed 1x with ice-
70
cold PBSCM (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, 1 mM CaCl2,
and 0.5 mM MgCl2, pH 7.4). The same number of cells from the two transfections were mixed
and fixed in 4 % paraformaldehyde for 15 min at room temperature in a 1.5 mL microcentrifuge
tube with gentle rotating. The fixed cells were permeabilized using PBSCM with 0.1 % Triton
X-100 for 15 min at room temperature, then blocked in PBSG (PBSCM with 40 % goat serum,
Gibco # 16210-064) for 30 min at room temperature with gentle rotating. For antibody staining,
these cells were incubated overnight at 4 oC in dark with gentle rotating with tau antibodies
diluted in PBSG (see Table 4.3 for dilution). The next day, the cells were incubated with either
Alexa Fluor 594 conjugated secondary anti-mouse IgG antibody (1:500, Invitrogen # A11005,
RRID: AB_2534073), anti-mouse IgM antibody (1:500, Invitrogen # A21044, RRID:
AB_2535713), or anti-rabbit IgG antibody (1:500, Invitrogen # A21442, RRID: AB_2535860)
diluted in PBSG for 1 hour at room temperature in dark with continuous rotating. Pelleting and
washing 1x with ice-cold PBSCM were required in between each incubation steps. After labeling,
the cells were re-suspended in ice-cold PBSCM, and the fluorescence signal from cells was
acquired using Becton-Dickinson Fortessa X20 (UConn Center for Open Research Resources
and Equipment). The experimenter was not blinded during the experiment.
To measure ΦSTT, the HEK293FT cells were co-transfected with plasmids pRK5-EGFP-tau WT
and pcDNA3-GSK3b. In separate wells, cells were transfected with pRK5-iRFP-tau WT plasmid
only (not co-transfected with pcDNA3-GSK3b), since these cells will be treated with λPP to de-
phosphorylate tau. The same number of cells from the two transfections were fixed and
permeabilized, separately. After permeabilization, only the cells expressing iRFP-tau WT were
treated with 2 U/µL λPP (New England Biolabs # P0753) for 3 hours at 30 oC with gentle
rotating. These cells were then mixed with cells expressing EGFP-tau WT. The following
71
blocking, primary and secondary antibody incubation steps follow the same procedure as when
measuring ΦNON, described above. The experimenter was not blinded during the experiment.
4.2.6 Immunocytochemistry in HEK293FT cells
The HEK293FT cells were seeded in a chamber mounted on #1.0 Borosilicate coverglass (Lab-
Tek # 155383), then co-transfected with plasmids pRK5-EGFP-tau WT and pcDNA3-GSK3b as
described above. Cells were rinsed 1x with ice-cold PBSCM and fixed in 4 % paraformaldehyde
for 15 min at room temperature in the chambers. The fixed cells were permeabilized using
PBSCM with 0.1 % Triton X-100 for 15 min at room temperature, blocked in PBSG for 30 min
at room temperature, and then incubated with tau antibodies diluted in PBSG overnight at 4 oC in
dark. The following tau antibodies and dilutions were used: AT270 (1:200), 1H6L6 (1:250), and
AT180 (1:200). The next day, the cells were incubated with either Alexa Fluor 594 conjugated
secondary anti-mouse antibody (1:500, Invitrogen # A11005, RRID: AB_2534073) or anti-rabbit
antibody (1:500, Invitrogen # A21442, RRID: AB_2535860) diluted in PBSG for 1 hour at room
temperature in dark. 300 nM of DAPI was used to stain the nucleus for 1 min at room
temperature. 3x washes with ice-cold PBSCM and 5 min incubation on ice in between were
required during each incubation steps. The cells were imaged using a Nikon A1R confocal
microscope (UConn Center for Open Research Resources and Equipment). The experimenter
was not blinded during the experiment.
4.2.7 Mouse hippocampus dissection
All procedures involving animals were in accordance with the US National Institutes of Health
Guide for the Care and Use of Laboratory Animals and approved by the University of
Connecticut Institutional Animal Care and Use Committee. Hippocampal dissection was
performed according to the methods we have previously described (2,3). No sample calculation
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was performed, but was based on previous studies (2,3). Timed pregnant Swiss Webster mice
(gestational day 15) were purchased from Taconic. P0 or P1 postnatal mice (randomly selected
from litter using simple randomization (4)) were placed on a physical barrier (such as a glove) on
top of an ice slurry one at a time to induce hypothermia to reduce or terminate movement and
bleeding, and decapitated. Hippocampus was dissected from 6 P0 or P1 postnatal mice in Hank’s
balanced salt solution with 10 mM HEPES, 35 mM glucose, 1 mM kynurenic acid, and 10 mM
MgCl2, pH 7.4. For dissociation, the hippocampal tissue was digested with 50 units of papain
(Worthington Biochemical) for 8 min, and halted with ovomucoid trypsin inhibitor (Worthington
Biochemical).
4.2.8 HEK293FT and mouse primary cell lysate preparation and western blotting
The following cell lysates were prepared: non-transfected HEK293FT cells, HEK293FT cells
expressing EGFP only, HEK293FT cells expressing EGFP-tau WT, and dissociated mouse
hippocampus. Cells were cultured to approximately 80 % confluency on 100 mm polystyrene
tissue culture plates and scraped into 1 mL lysis buffer containing 150 mM NaCl, 5 mM EDTA,
50 mM Tris, 1 % Triton X-100, 0.5 % sodium deoxycholate, and 0.1 % sodium dodecyl sulfate,
protease inhibitor cocktail (Sigma # P8340), and phosphatase inhibitor cocktails 2&3 (Sigma #
5726 & P0044). Cells were lysed on ice for 15 min and then centrifuged at 13,000 g for 15 min
at 4 oC to remove insoluble debris. The concentrations of cell lysate were determined using a
Bradford assay kit (ThermoFisher Scientific # 23200) according to the manufacturer’s
instructions. The lysates were aliquoted and stored at -20 oC avoiding multiple freeze/thaw
cycles.
An equal amount (15 µg) of total cellular protein was separated by SDS-PAGE using 4-12.5 %
Bis-Tris gels and transferred to nitrocellulose membrane (GE Healthcare # 10600006). SeeBlue
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pre-stained protein standard (Invitrogen # LC5925) was used to determine the molecular weight.
The nitrocellulose membrane was blocked in TBST (20 mM Tris, 150 mM NaCl, and 0.1 %
Tween 20, pH 7.6) with 1 % BSA (bovine serum albumin, ThermoFisher Scientific #149006) for
1 hour at room temperature, and then incubated with the tau antibodies diluted in TBST with 1 %
BSA overnight at 4 oC with continuous agitation. The following antibodies and dilutions were
used: AT270 (1:1000), 1H6L6 (1:2500), and AT180 (1:1000). The following day, the membrane
was washed 3x with TBST and then incubated with either horseradish peroxidase-coupled
secondary anti-mouse antibody (1:2000, Invitrogen # 62-6520, RRID: AB_2533947) or anti-
rabbit antibody (1:5000, Invitrogen # 31460, RRID: AB_228341) for 1 hour at room temperature
with continuous agitation. The peroxidase activity was revealed using an enhanced
chemiluminescence detection kit (GE Healthcare, # 16915806). GAPDH antibody (1:2000,
Invitrogen # MA5-15738-HRP, RRID: AB_2537659) was used as a loading control. The
experimenter was not blinded during the experiment.
Results
4.3.1 𝜱: A generalizable parameter for measuring the specificity of phospho-tau antibodies
To quantify the specificity of phospho-tau antibodies, we developed a whole-cell based approach
using flow cytometry (Figure 4.1). To probe the site-specificity of phospho-tau antibodies, we
generated targeted alanine mutants at defined phosphorylation sites. HEK293FT cells were
transiently transfected with either WT human tau (0N4R isoform) fused to the EGFP (EGFP-tau
WT), or a point mutant (an alanine mutant at the phosphorylation site of interest) fused to the
iRFP (iRFP-tau), in separate wells (Figure 4.1a). Previous reports indicated that endogenous tau
expression is undetected in HEK293 cells (5,6). In addition, when transfected with human tau,
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endogenous kinases in HEK293FT cells phosphorylated human tau (7,8). Leveraging these facts,
we developed a ratiometric assay to quantify the specificity of phospho-tau antibodies (Figure
4.1a). The EGFP-positive cells (expressing WT tau) and iRFP-positive cells (expressing a tau
point mutant incapable of phosphorylation at a defined site) were mixed in same numbers and
incubated with phospho-tau antibodies (Figure 4.1a). The EGFP (green channel) and iRFP (far-
red channel) positive cells, and their antibody binding (red channel) were clearly distinguished
using flow cytometry (Figure 4.1b, left panel). This assay quantifies the specific binding of the
antibody to a defined phosphorylation site. In this assay, the phospho-tau antibody AT180,
targeting phospho-threonine 231 (pThr231) (9-11), showed almost exclusive labeling to the cells
expressing EGFP-tau WT, and no detectable binding to iRFP-tau T231A (Figure 4.1b),
indicating that AT180 binding requires pThr231.
Figure 4.1 A ratiometric assay to quantify the specificity of phospho-tau antibodies.
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a, schematic of phospho-tau antibody specificity quantification assay using HEK293FT. HEK293FT cells were transfected with either EGFP-tau WT (12) or iRFP-tau with alanine mutation(s) at the target site (magenta), in separate wells. The cells were then mixed and incubated with phospho-tau antibodies. The antibody binding was detected using an Alexa Flour 594 conjugated secondary antibody (red). In parallel, HEK293FT cells were transfected with either EGFP-tau WT (12) or iRFP-tau WT (magenta), in separate wells. iRFP-tau WT cells were treated with lambda phosphatase (λPP) to de-phosphorylate tau, then mixed with EGFP-tau WT cells. Scale bars, 50 µm. b, representative flow cytometry results for AT180 showing the alanine mutant experiment (left) and de-phosphorylation experiment (right).
In case an antibody shows non-specific binding to the alanine mutant at the target modification
site, the antibody may be recognizing a phosphorylated residue at an off-target site. In addition,
the absence of antibody binding to the alanine mutant does not exclude the possibility that
binding to the non-modified tau is inhibited by the alanine mutation. In other words, the antibody
may possess non-specific binding to unmodified tau, but may not bind to the alanine point
mutant. To assess these possibilities, we transfected HEK293FT cells with either EGFP-tau WT
or iRFP-tau WT separately and treated only the iRFP-tau WT expressing cells with λPP (Figure
4.1a). As in the assay using the T231A mutant, AT180 showed exclusive binding to EGFP-tau
WT cells (Figure 4.1b, right panel). This result provides further support that the AT180 antibody
does not recognize the WT tau epitope without phosphorylation of Thr231.
We tested the assay (Figure 4.1a) for its ability to discern phospho-specificity using 3 previously
characterized anti-tau antibodies. Tau-5 is a pan-tau antibody that shows phosphorylation-
independent binding (13,14), 3.24 is a high specificity scFv to pThr231 tau (15), and ab131354
targets pSer262 tau but have shown cross-reactivity to non-phosphorylated tau (16). Tau-5
indeed showed binding to both EGFP-tau WT and iRFP-tau WT treated with λPP (Figure 4.2a).
3.24 showed binding to EGFP-tau WT, but not to iRFP-tau T231A or iRFP-tau WT treated with
λPP (Figure 4.2b), confirming its high specificity. ab131354 showed cross-reactivity to both
iRFP-tau S262A and iRFP-tau WT treated with λPP (Figure 4.2c), indicating that it is not
specific to pSer262 tau.
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Figure 4.2 Validation of the assay using well characterized anti-tau antibodies a, flow cytometry plots for the pan-tau antibody Tau-5, which shows binding to both EGFP-tau WT and iRFP-tau WT treated with λPP. b-c, flow cytometry plots for the high specificity single-chain variable fragment 3.24 (b) and ab131354 (c).
Based on this assay, we introduce a parameter termed Φ (see Materials and Methods section for
definition) that quantifies the specificity of phospho-site targeting antibodies. Φ is determined by
the degree of cross-reactivity towards cells expressing the target antigen with an alanine point
mutation at the target phospho-site, or cells expressing the target antigen but were treated with
λPP for de-phosphorylation. Φ ranges from 0 to 1, where 1 indicates perfect specificity (no
binding to the alanine mutant or λPP treated cells), and lower Φ indicates more false positive
detection of nonphospho-tau or phospho-tau at off-target sites. 2 Φ values are derived from the
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definition: ΦVWV is determined from cross-reactivity to tau with a point alanine mutation at the
target site, while ΦSTT is determined by cross-reactivity to de-phosphorylated tau. The phospho-
site specificity is determined by evaluating both ΦVWV and ΦSTT values. The specificity ΦVWV and
ΦSTT of AT180 were 0.99 ± 0.01 and 0.98 ± 0.01, respectively (Table 4.3, mean ± standard
deviation from 2 independent experiments, all data points shown in Figure 4.3), indicating
nearly perfect phospho-site specificity towards pThr231 tau. The specificity ΦVWV and ΦSTT of
ab131354 were 0.32 ± 0.02 and 0.37 ± 0.01, respectively, reflecting its poor specificity (Figure
4.2). Φ can be used as a general standard to measure and compare the specificity of post-
translational modification (PTM)-site targeting antibodies.
Figure 4.3 Measurement of 𝚽𝐚𝐥𝐚 and 𝚽𝛌𝐏𝐏 of phospho-tau antibodies. Error bars indicate standard deviation from two independent cell culture preparations. Red, ΦSTT. Blue, ΦVWV.
4.3.2 Co-expression of glycogen synthase kinase-3b enhances tau phosphorylation
The assay developed here relies on intracellular phosphorylation of tau due to endogenous kinase
activity in HEK293FT cells. Previous studies have found that tau expressed in HEK293 cells
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were phosphorylated at various sites, including S214, S262, S396/S404 (17). A major advantage
of our assay is the potential to co-express kinases to induce tau phosphorylation. Since many of
the residues that are phosphorylated on tau are serine or threonine preceding a proline, we co-
expressed glycogen synthase kinase (GSK)-3b, a proline-directed kinase, in HEK293FT cells to
enhance tau phosphorylation. GSK-3b phosphorylates tau at a wide range of sites found in
Alzheimer’s disease patients (18). Co-transfected cells showed a similar level of EGFP-tau WT
expression (Figure 4.4a), but enhanced binding to the phospho-tau antibody AT8, which targets
pSer202 (19-21) (Figure 4.4b), indicating improved phosphorylation. Therefore, to enhance the
detection of phospho-tau antibody binding, we co-transfected tau constructs with GSK-3b in
subsequent experiments.
Figure 4.4 Co-expression of GSK-3b in HEK293FT cells enhances phosphorylation of tau. a, representative flow cytometry plots for AT8 showing tau expression (EGFP-tau) and AT8 binding cells transfected with tau-only (left) or cells co-transfected with tau and GSK-3b (right). b, histograms for tau expression (left) and AT8 binding (right) in cells transfected with tau-only (red) or cells co-transfected with tau and GSK-3b (blue).
4.3.3 Quantifying the phospho-site specificity of commonly used phospho-tau antibodies
Using our assay, we quantified the specificity Φ of 7 widely used phospho-tau antibodies
(AT270, AT8, AT100, AT180, PHF-6, TG-3, and PHF-1) as well as 2 additional phospho-tau
antibodies (1H6L6 and pS404). Collectively these antibodies have been raised against 6 distinct
phosphorylation sites (see Table 4.3 for more information) of high relevance in a wide range of
79
neurodegeneration. We applied the same approach described in Figure 4. 1 to quantify both ΦVWV
and ΦSTT. In Figure 4.5, data in the left two columns show antibody labeling from a mixture of
cells transfected with either EGFP-tau WT or iRFP-tau with the corresponding alanine point
mutant. The right two columns show antibody labeling from cells transfected with either EGFP-
tau WT or iRFP-tau WT treated with λPP. Clear binding to EGFP-tau WT was detected in all
antibodies except AT100 (Figure 4.5, leftmost column). Most antibodies showed exclusive
binding to EGFP-tau WT, and no binding to the alanine point mutant at their corresponding
target site or the iRFP-tau WT treated with λPP (Figure 4.5, second and last columns from left).
From two independent experiments, we obtained highly reproducible measurement of ΦVWV and
ΦSTT from these antibodies (Table 4.3 and Figure 4.3). 5 out of 7 antibodies (AT8, AT180,
PHF-6, TG-3, and PHF-1) had Φ values near 1, indicating these antibodies have nearly perfect
specificity. For AT270, ΦVWV = 0.80 ± 0.06 (Table 4.3), due to the elevated level of binding
signal in cells expressing the T181A mutant (Figure 4.5). However, ΦSTT = 0.99 ± 0.01 (Table
4.3), indicating that the cross-reactivity was not detected in dephosphorylated cells. Taken ΦVWV
and ΦSTT together, the non-specific signal detected from AT270 was phosphorylation-sensitive
(see below for further characterization of non-specific binding in AT270). For AT100, the
absence of binding to EGFP-tau WT cells suggests that the phospho-site (pThr212/pSer214) was
not phosphorylated in HEK293FT cells, even with co-transfection of GSK-3b. Therefore, we
were unable to measure the Φ values for AT100. Taken together, these results validate that the
widely used phospho-tau antibodies AT8, AT180, PHF-6, TG-3, and PHF1 binds specifically to
tau phosphorylated at the target site, but not to non-phosphorylated tau or phosphorylated tau at
off-target sites.
80
81
Figure 4.5 Specificity measurements for commonly used phospho-tau antibodies Specificity measurements for commonly used phospho-tau antibodies using the flow cytometry assay described in Figure 4.1. Dotted and closed circles indicate observed non-specific binding to cells not expressing tau.
4.3.4 Non-specific binding to cells in phospho-tau antibodies
For the antibody AT270, which showed lower ΦVWV but high ΦSTT, we observed a reproducible
non-specific binding to cells not expressing tau (dotted circles in Figure 4.5). Similar non-
specific binding was detected with AT100, even though AT100 did not bind to EGFP-tau WT.
Although not as prominent, a small fraction and weak level of non-specific binding to tau-
negative cells was detected in antibodies 1H6L6 and TG-3 (closed circles in Figure 4.5). For
other antibodies, background binding to non-transfected cells was not detected. The binding to
cells not expressing tau indicates that these antibodies exhibit non-specific binding to
HEK293FT cells. We conducted experiments to rule out the possibility that the non-specific
binding is originating from the secondary reagents used for detection. Control antibodies having
matched isotype (mouse IgG1, mouse IgM, or rabbit IgG) followed by the secondary antibodies
(see Materials and Methods section) were used to label a mixture of cells transfected with either
EGFP-tau WT, or cells transfected with iRFP-tau WT treated with λPP (Figure 4.6). No non-
specific binding was detected in the secondary reagents, indicating that the non-specific binding
observed was not due to secondary reagents.
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Figure 4.6 Flow cytometry plots showing binding of isotype-matching control antibodies under the specificity quantification assay conditions
For AT270 and AT100, the population with non-specific binding separated clearly when cells
were treated with λPP (Figure 4.5, right two columns). Based on this result, we hypothesized
that the non-specific bindings of AT270 and AT100 are due to cross-reactivity to other
phosphorylated proteins in HEK293FT cells. In this scenario, cells treated with λPP would show
decreased level of antibody binding, while untreated cells would have generally elevated
antibody binding. To test this, we labeled a mixture of cells transfected with either EGFP (not
EGFP-tau), or cells transfected with iRFP (not iRFP-tau) treated with λPP. In this experiment,
binding was detected only in GFP-positive cells (Figure 4.7a). Since specific binding to
phospho-tau in AT100 was not detected, we further analyzed the non-specific binding of AT270.
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We also analyzed the non-specific binding of 1H6L6, which showed a weak level of non-specific
binding (closed circles in Figure 4.5). TG-3 was not included in this analysis since the fraction
of tau-negative cells showing non-specific binding was even smaller. To further assess the non-
specific binding found in AT270 and 1H6L6, we conducted immunocytochemistry experiments.
Antibodies AT270 and 1H6L6 showed binding in cells not expressing EGFP-tau, while the
highly specific antibody AT180 showed staining only to EGFP-tau positive cells (Figure 4.7b).
To assess the broadness of proteins that cause the non-specific binding, we conducted western
blotting experiments. We probed cell lysates from non-transfected, EGFP-transfected, EGFP-tau
transfected cells as well as dissociated mouse hippocampus. In these experiments, antibodies
AT270 and 1H6L6 showed bands in addition to a band corresponding to EGFP-tau (Figure 4.7c).
In contrast, a single band corresponding to EGFP-tau was observed in AT180 (Figure 4.7c). For
AT270, off-target bands found in HEK293FT cell lysates were also detected in WT mouse
hippocampal cell lysate. Therefore, this non-specific binding was independent of species, leading
to a non-specific band interfering with the mouse tau signal. However, for 1H6L6, the non-
specific binding was not clearly detected in mouse hippocampal lysate, suggesting that the non-
specific binding may be species or cell-type dependent. These results demonstrate that by using a
whole-cell based approach, we have identified a source of non-specific binding in antibodies
AT270 and 1H6L6 not observable when using peptides for specificity characterization.
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Figure 4.7 Non-specific binding to HEK293FT cells in phospho-tau antibodies a, flow cytometry plots for AT270 and AT100 to detect non-specific binding to normal HEK293FT cells (EGFP-transfected) vs. λPP-treated cells (iRFP-transfected). b, confocal images of AT180, AT270, and 1H6L6 antibodies binding to HEK293FT cells transfected with EGFP-tau WT. Images show nuclei stained using DAPI (blue), tau expression (12), antibody staining (red), and merge. Scale bars, 25 µm. c, analysis of tau signal and non-specific signal in AT270 and 1H6L6 by western blotting. The proteins were extracted from: lane 1, non-transfected HEK293FT cells; lane 2, HEK293FT cells transfected with EGFP; lane 3, HEK293FT cells co-transfected with EGFP-Tau/GSK-3b; lane 4, mouse hippocampus. Proteins were identified with the following antibodies: AT180, AT270 , and 1H6L6. GAPDH was used as a loading control.
Discussion
Site-specific phosphorylation of tau is critical in understanding the molecular biology of tau and
is a well-defined target in neurodegeneration. The specificity of phospho-tau antibodies is critical
for reproducible and sensitive detection (15,22). Phospho-tau antibodies should interact only
with the target epitope sequence with site-specific phosphorylation, and not with nonphospho-tau
85
or phospho-tau at off-target sites. In particular, considering the fact that phospho-tau antibodies
are being developed for diagnostic tests and clinical trials as therapeutics, a standard measure of
specificity is urgently needed (23-28) In this study, we developed a whole-cell based assay that
enables robust measurement of the specificity of phospho-tau antibodies, and introduced the
specificity parameter termed Φ. Φ is determined by measuring the fraction of binding signal
from cross-reactivity to nonphospho-tau or off-target phosphorylation sites. Φ ranges from 0 to 1,
where 1 indicates perfect specificity and lower Φ indicates more false positive detection of either
nonphospho-tau sequences or the phospho-tau at the off-target site. We measured two Φ values
(ΦVWV and ΦSTT ) to comprehensively capture the specificity. ΦVWV indicates the phospho-site
specificity of the antibody by measuring the fraction of non-specific binding to an alanine point
mutant at the target phospho-site. ΦSTT indicates the phospho-sensitivity of the antibody by
measuring non-specific binding to de-phosphorylated tau. This approach can be easily adapted to
other phosphorylated proteins, and potentially to other PTMs such as methylation and acetylation,
allowing specificity quantification and comparison of any PTM-site specific antibody. In
addition, the Φ values can be combined to a single parameter, to yield a single specificity
parameter. For example,
ΦXYXVW = 1 − ((1 − ΦVWV) + (1 − ΦSTT))
= ΦVWV + ΦSTT − 1
In our calculation of the Φ values, we did not include the fluorescence signal detected from cells
not expressing tau. Therefore, the small non-specific binding to cells we observed for antibody
1H6L6 (and to an even smaller degree in TG-3) is not reflected in their Φ values. As for AT270,
the Φ value captured the non-specific binding to cells, since the non-specific binding was
detected in the cells expressing the alanine mutant but was lost after de-phosphorylation with
86
λPP. We attempted to define another Φ parameter to quantify the non-specific binding to cells
but the resulting values were inconsistent due to variations in the level of non-specific binding
signal and fraction of cells showing binding.
Using our assay, we measured the Φ values of 10 phospho-tau antibodies, 7 of which have been
used in hundreds of previous studies. Out of the 7, we quantified the specificity of 6 antibodies
(AT270, AT8, AT180, PHF-6, TG-3, and PHF-1). We were unable to determine Φ values for
AT100, due to lack of binding to tau expressed in HEK293FT cells. Endogenous kinases in
HEK293FT cells phosphorylate human tau (7,8). We showed that co-transfection of tau with
GSK-3b led to increased phosphorylation of tau, but not for the AT100 target site
(Thr212/Ser214). Previous studies have indicated that the AT100 epitope is formed only when a
specific order of phosphorylation occurs (29). It has also been shown that Thr212 and Ser214 are
sequentially phosphorylated by GSK-3b and protein kinase A (PKA), respectively (29).
Considering that PKA is commonly expressed in cells, tau expressed in HEK293FT cells may
only be phosphorylated at Ser214, leading to lack of AT100 binding. Tau phosphorylation may
be enhanced by using mammalian brain extracts, which contain kinases that phosphorylate tau in
vitro (29-31).
Out of the 6 phospho-tau antibodies (AT270, AT8, AT180, PHF-6, TG-3, and PHF-1), all but
AT270 had Φ values close to 1, indicating near-perfect specificity. For AT270, a weak, but
reproducible binding was detected to tau-negative HEK293FT cells (Figure 4.5). This non-
specific binding was consistently detected in immunocytochemistry experiments, leading to
distinct perinuclear non-specific binding (Figure 4.7b). Western blotting with AT270 revealed
additional bands consistently present in non-transfected HEK293FT cell lysate as well as
primary WT mouse hippocampal lysate (Figure 4.7c). A previous study found multiple bands
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when probing WT mouse brain lysate (32). The same study also found non-specific binding in
antibodies AT8, AT180, and TG-3, using cell lysate prepared from tau knockout mice. However,
this non-specific binding was not due to the antibodies themselves, and was eliminated by
switching secondary reagents (32). We found that the two Φ values for AT270 was inconsistent
(ΦVWV = 0.80 ± 0.06 vs. ΦSTT = 0.99 ± 0.01). This indicated that the source of cross-reactivity
was lost upon de-phosphorylation with λPP, suggesting that AT270 may bind to another
phosphorylated protein. Indeed, the non-specific binding was only detected from the tau-negative
cells expressing EGFP, but not from the tau-negative cells expressing iRFP with de-
phosphorylation treatment (Figure 4.7a).
A major advantage of our specificity assay is the use of flow cytometry, which allows sensitive,
quantitative detection of antibody binding. We enabled the distinction of non-specific binding
within a single sample by tagging potential cross-reactive proteins with a fluorescent protein
easily distinguishable from the WT target protein. These features led to the reproducible
measurement of Φ values for a wide range of phospho-tau antibodies. Previously, peptide arrays
have been used to validate the specificity of PTM-specific antibodies, such as those targeting
histone modifications (33-35). Synthetic peptides and peptide arrays were also used for
specificity analysis of phospho-tau antibodies (16). The limitation of this approach is the high
cost of synthesizing peptides in order to comprehensively cover tau phosphorylation sites.
Moreover, short peptides do not have strong binding to conformation-sensitive phospho-tau
antibodies, such as TG-3 (10). Our approach allows assessment of binding to full-length tau with
defined sequence variations, and can be co-expressed with kinases to induce tau phosphorylation.
Using our approach, the specificity of the conformation-sensitive pT231 tau antibody TG-3 was
also successfully determined (Figure 4.5 and Table 4.3). Libraries of tau mutations may be
88
generated, and deep sequencing approaches can be used for de novo identification of anti-tau
antibodies. Our approach also revealed non-specific binding of antibodies to irrelevant cellular
proteins. We found that the source of non-specific binding for AT270 is conserved across
HEK293FT cells and mouse primary hippocampal cells, but was not conserved for 1H6L6,
indicating cell-type and species sensitivity of non-specific binding. Further analyses using
immunoprecipitation and mass spectrometry may allow the identification of the cross-reactive
proteins.
Since the cross-reactivity of antibodies would be dependent on the affinity to the off-target sites
or molecules, it is possible that the Φ values are dependent on antibody concentration. In
particular, the Φ values may decrease upon increasing antibody concentration. In our
experiments, we used antibody concentrations typically used for cell labeling experiments (2
µg/mL, Table 4.3). Since we observed high antibody binding signal in these experiments for all
antibodies tested except AT100 (which showed non-specific binding to cells), we did not attempt
to increase the antibody concentration. Also, all antibodies that showed binding had near-perfect
specificity, except AT270 and ab131354. Lowering the antibody concentration for these
antibodies resulted in a significant decrease in antibody binding, and did not result in
improvement of Φ values (data not shown). Even for the high specificity antibodies, users should
be cautious when using them at higher concentrations than what is listed in Table 4.3, since that
may result in increased non-specific binding.
Conclusion
In summary, our study provided a whole-cell based approach to validate the specificity of
phospho-tau antibodies. Using this assay, we determined the parameter Φ for phospho-tau
89
antibodies, that enable quantitative comparison of specificity. We anticipate that this parameter
will be widely adopted in guiding users of the specificity of phospho-tau antibodies, and
potentially PTM-specific antibodies in general.
90
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16. Ercan, E., Eid, S., Weber, C., Kowalski, A., Bichmann, M., Behrendt, A., Matthes, F., Krauss, S., Reinhardt, P., Fulle, S., and Ehrnhoefer, D. E. (2017) A validated antibody panel for the characterization of tau post-translational modifications. Mol Neurodegener 12, 87
17. Ren, Q. G., Liao, X. M., Chen, X. Q., Liu, G. P., and Wang, J. Z. (2007) Effects of tau phosphorylation on proteasome activity. FEBS Lett 581, 1521-1528
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18. Mandelkow, E. M., Drewes, G., Biernat, J., Gustke, N., Van Lint, J., Vandenheede, J. R., and Mandelkow, E. (1992) Glycogen synthase kinase-3 and the Alzheimer-like state of microtubule-associated protein tau. FEBS Lett 314, 315-321
19. Pei, J. J., Braak, E., Braak, H., Grundke-Iqbal, I., Iqbal, K., Winblad, B., and Cowburn, R. F. (1999) Distribution of active glycogen synthase kinase 3beta (GSK-3beta) in brains staged for Alzheimer disease neurofibrillary changes. J Neuropathol Exp Neurol 58, 1010-1019
20. Sereno, L., Coma, M., Rodriguez, M., Sanchez-Ferrer, P., Sanchez, M. B., Gich, I., Agullo, J. M., Perez, M., Avila, J., Guardia-Laguarta, C., Clarimon, J., Lleo, A., and Gomez-Isla, T. (2009) A novel GSK-3beta inhibitor reduces Alzheimer's pathology and rescues neuronal loss in vivo. Neurobiol Dis 35, 359-367
21. Hernandez, F., Lucas, J. J., Cuadros, R., and Avila, J. (2003) GSK-3 dependent phosphoepitopes recognized by PHF-1 and AT-8 antibodies are present in different tau isoforms. Neurobiol Aging 24, 1087-1094
22. Lothrop, A. P., Torres, M. P., and Fuchs, S. M. (2013) Deciphering post-translational modification codes. FEBS Lett 587, 1247-1257
23. Chai, X., Wu, S., Murray, T. K., Kinley, R., Cella, C. V., Sims, H., Buckner, N., Hanmer, J., Davies, P., O'Neill, M. J., Hutton, M. L., and Citron, M. (2011) Passive immunization with anti-Tau antibodies in two transgenic models: reduction of Tau pathology and delay of disease progression. J Biol Chem 286, 34457-34467
24. Boutajangout, A., Ingadottir, J., Davies, P., and Sigurdsson, E. M. (2011) Passive immunization targeting pathological phospho-tau protein in a mouse model reduces functional decline and clears tau aggregates from the brain. Journal of neurochemistry 118, 658-667
25. Kayed, R. (2010) Anti-tau oligomers passive vaccination for the treatment of Alzheimer disease. Hum Vaccin 6, 931-935
26. Asuni, A. A., Boutajangout, A., Quartermain, D., and Sigurdsson, E. M. (2007) Immunotherapy targeting pathological tau conformers in a tangle mouse model reduces brain pathology with associated functional improvements. J Neurosci 27, 9115-9129
27. Boutajangout, A., Quartermain, D., and Sigurdsson, E. M. (2010) Immunotherapy targeting pathological tau prevents cognitive decline in a new tangle mouse model. J Neurosci 30, 16559-16566
28. Lewis, J., McGowan, E., Rockwood, J., Melrose, H., Nacharaju, P., Van Slegtenhorst, M., Gwinn-Hardy, K., Paul Murphy, M., Baker, M., Yu, X., Duff, K., Hardy, J., Corral, A., Lin, W. L., Yen, S. H., Dickson, D. W., Davies, P., and Hutton, M. (2000) Neurofibrillary tangles, amyotrophy and progressive motor disturbance in mice expressing mutant (P301L) tau protein. Nat Genet 25, 402-405
29. Zheng-Fischhofer, Q., Biernat, J., Mandelkow, E. M., Illenberger, S., Godemann, R., and Mandelkow, E. (1998) Sequential phosphorylation of Tau by glycogen synthase kinase-3beta and protein kinase A at Thr212 and Ser214 generates the Alzheimer-specific epitope of antibody AT100 and requires a paired-helical-filament-like conformation. Eur J Biochem 252, 542-552
30. Gustke, N., Steiner, B., Mandelkow, E. M., Biernat, J., Meyer, H. E., Goedert, M., and Mandelkow, E. (1992) The Alzheimer-like phosphorylation of tau protein reduces microtubule binding and involves Ser-Pro and Thr-Pro motifs. FEBS Lett 307, 199-205
31. Lichtenberg-Kraag, B., Mandelkow, E. M., Biernat, J., Steiner, B., Schroter, C., Gustke, N., Meyer, H. E., and Mandelkow, E. (1992) Phosphorylation-dependent epitopes of neurofilament antibodies on tau protein and relationship with Alzheimer tau. Proc Natl Acad Sci U S A 89, 5384-5388
32. Petry, F. R., Pelletier, J., Bretteville, A., Morin, F., Calon, F., Hebert, S. S., Whittington, R. A., and Planel, E. (2014) Specificity of anti-tau antibodies when analyzing mice models of Alzheimer's disease: problems and solutions. PloS one 9, e94251
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Conclusion
This work has addressed the problems of generating and characterizing high-specific and high-
affinity phospho-tau antibodies. We introduced a yeast display approach to quantify the
specificity of PTM-specific antibodies, and developed a screening assay based on it to generate
PTM-specific antibodies with improved affinity and high specificity. We demonstrated that
improving PTM-specific antibody affinity through directed evolution results in cross-reactivity
to the non-modified target sequence, there’s a trade-off between affinity and specificity in PTM-
specific antibodies. The results from this study demonstrate that in vitro affinity maturation is a
viable option for improving the affinity of PTM-specific antibodies. In this process, it is essential
to quantify the specificity, particularly to the non-modified target sequence. By quantifying the
specificity, it is possible to apply selection pressures for improved affinity and specificity
simultaneously.
We also developed a whole-cell based approach to quantify the specificity of phospho-tau
antibodies and introduced a parameter Φ to measure their specificity. We anticipate that this
parameter will be widely adopted in guiding the use of phospho-tau antibodies, and PTM-
specific antibodies in general.
Future perspectives
5.2.1 Avidity effect on antibody screening assay
Antibody affinity and avidity
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Antibody affinity measures the interaction strength between an epitope and the antigen binding
site on antibody. It is defined by a measure of dynamic equilibrium of the reversible
biomolecular interaction.
Calculation of binding affinity for antibody-antigen interaction (1 antibody binding site per 1
antigen):
𝐴𝑏 + 𝐴𝑔 ⟺ [𝐴𝑏 − 𝐴𝑔]
where [Ab] is the unbound antibody concentration, [Ag] is the unbound antigen concentration,
and [Ab-Ag] is the concentration of the antibody-antigen complex.
Calculation of dissociation constant, which is the affinity constant (Kd):
𝐾𝑑 =𝑘E``𝑘E8
= 𝐴𝑏 [𝐴𝑔][𝐴𝑏 − 𝐴𝑔]
where 𝑘E8 is a constant used to characterize how quickly the antibody binds to its target, and
𝑘E`` is a constant used to characterize how quickly an antibody dissociates from its target.
High affinity antibodies bind quickly to the antigen, allowing sensitive binding and detection in
assays under difficult conditions. Low affinity antibodies bind slowly and weakly to the antigen
and often fail to detect the antigen in assays.
Antibody avidity refers to the accumulated strength of multiple antigen-binding sites interact
with the target epitopes simultaneously. Avidity is thought as the combined effect of all affinities
involved in the biomolecular interaction, which is referred to as functional affinity (1). Nature
uses multivalency to govern many biological processes.
Avidity effect on antibody screening assay
All antibodies are at least bivalent or multivalent, however, the engineered binding proteins
developed from in vitro screening assay are generally monovalent. When generating novel scFvs
or nanobodies using in vitro screening assay, it is extremely difficult to discover binders in the
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initial rounds of screening due to their low affinity. The greater valency of an antibody, the
greater the amount of antigen it can bind. Similarly, antigens can demonstrate multivalency
because they can bind to more than one antibody. Multimeric interactions between an antibody
and an antigen slow their dissociation and stabilize the binding. Individually, each binding
interaction may dissociate quickly, however, when many binding interactions are present at the
same time, transient unbinding of a single site does not allow the molecule to dissociate away,
and binding of that weak interaction is likely to be restored. Therefore, we hypothesize that
avidity effect on yeast displayed scFvs can increase the possibility to obtain rare low affinity
binders, and we can develop a new screening assay based on it.
In order to test our hypothesis, we will assess the impact of avidity of yeast displayed pT231
scFv on its binding to the target phosphopeptide. Three parameters are involved in this study,
and the relevant specific aims are:
i. Affinity of the scFv for the phosphopeptide
In Chapter 3, we have identified some scFv mutants those only bind to target phosphopeptide
but have no cross-reactivity to nonphospho-peptide or scrambled phosphopeptide. We will assess
the affinity of yeast displayed mutants for the target phosphopeptide and select the high-affinity
mutants, moderate-affinity mutants, and low-affinity mutants. We will measure the dissociation
rate of these mutants.
ii. Valency of the phosphopeptide
The valency of the phosphopeptide can be increased by streptavidin-biotin interaction.
Streptavidin has the ability to bind up to four biotin molecules (Figure 5.1) with an extremely
high affinity (Kd=10-14 M) (2). The bond between streptavidin and biotin is one of the strongest
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non-covalent interaction. It is unaffected by pH, temperature, organic solvents and other
denaturing reagents once formed.
Figure 5.1 Schematic of the Streptavidin-biotin interaction Streptavidin can bind up to four biotin or biotinylated molecules, which is conjugated to a target phosphopeptide to form a streptavidin-biotin complex.
The valency of phosphopeptide can be controlled by incubating the streptavidin with different
ratios of biotinylated phosphopeptide and free biotin. We will label the yeast displayed scFv
mutants with those pre-incubated phosphopeptide complex, and measure the dissociation rate of
each mutants in different conditions.
iii. Structural arrangement of the functional scFv
More functional scFvs expression per yeast cell also contributes to the avidity effect. A favorable
structural arrangement of functional scFvs increases the local concentration of phosphopeptide
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binding sites. Once the first phosphopeptide binds to an scFv on the yeast surface, the chance of
a bivalent, trivalent, and/or tetravalent interaction are improved.
To enable this idea, we will assess the impact of linker length between yeast wall protein Aga2p
and scFv on avidity effect (Figure 5.2). We will insert a longer polypeptide linker between
Aga2p and the pT231 scFv, allowing more functional scFvs cluster together. We will measure
the dissociation rate of the same scFv in different yeast surface display formats.
Figure 5.2 Schematic of yeast surface display formats The yeast surface display vector encodes scFv as a fusion to the C-terminus of Aga2p with a shorter polypeptide linker (left), and a longer polypeptide linker (right).
5.2.2 Application of binding proteins in optogenetics
The discovery of light-gated ion channels and their application to control neural activities have
had a transformative impact on the field of neuroscience. In recent years, the concept of using
light-activated proteins to control biological processes has greatly diversified into other fields,
driven by the natural diversity of photoreceptors and decades of knowledge obtained through
their biophysical characterization. This strategy, commonly referred to as optogenetics, is
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particularly powerful in deciphering the role of targeted neural cells within their complex native
environment.
The development intracellular binding proteins for neural proteins, such as tau, combined with
optogenetics tools allows the visualization of complex structures such as the synapse that are
critical for understanding neural function. These technologies have the potential to pinpoint the
location of proteins in fine neuronal structures and reveal the changes occurring in these
structures under diseased state. Therefore, our screening assay on improved affinity and high
specific antibody will be applied to develop intracellular binding proteins for key neural target
proteins and coupling optogenetics to these binding proteins for studying target protein function
in the future.
Introduction
The word ‘optogenetics’ was initially used within the context of neuroscience (3), to describe the
approach of using light to image and control neuronal activity in the intact, living brain. The idea
of controlling biological function using light has been around for a long time, but it is in the field
of neuroscience that its need as a truly enabling tool has been envisioned early on (4,5), and
remarkably to the realization of such potential (6-8) with wide acceptance over the past decade.
The success of optogenetics in neuroscience has sparked interest of many scientists and
engineers in other fields, and now the definition of optogenetics has broadened to encompass the
general field of biotechnology that combines genetic engineering and optics to enable gain or
loss of well-defined function, often in the intact animal (9,10).
Original concepts: a brief historical perspective
The seed for the birth of optogenetics in neuroscience may have been planted for a long time by
the pioneering work of Ramón y Cajal, who provided foundational evidence that neurons are the
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signaling units of the nervous system, and that they exist in many distinctive morphologies (11).
Since then, studies followed to show that indeed many different types of neurons exist, classified
based on their physiological characteristics, anatomical location, morphology, and gene
expression profile (12-14). It is still unclear how many cell types exist in the human brain, but it
is speculated that there are about 1,000 neuronal cell types just within the mammalian cortex
(15,16). In a general sense, it is agreed that a specific cell type carries out the same function
within a neural circuit (17). Therefore, identification of all cell types and mapping out their
connectivity is essential in order to understand how the nervous system works. Identification of
different cell types must be accompanied with functional characterization within their normal
context, the nervous system. For this reason, neuroscientists have been looking for a way to
perturb individual cell types within the intact brain. This was clearly expressed by Francis Crick
in his insightful discussion published in 1979, in which he proposed the need for a ‘method by
which all neurons of just one type could be inactivated, leaving the others more or less unaltered’
(4).
Optogenetics is arguably the first technological breakthrough that enables such experiments. It
makes the crucial connection between cell-type information and the ability to perform gain or
loss of function experiments. Prior to the development of optogenetics, experimental approaches
existed in neuroscience to use light as a way to control neural activity. For example, ‘caged’
compounds such as secondary messenger molecules, ions, and neurotransmitters have been
developed that are initially inert, but become active upon light illumination (18,19). Even
though these photochemical approaches did not provide ways to control specific cell-types, they
laid the groundwork for the use of millisecond timescale illumination in intact cells and tissue
(19). Cell-type specific activation of neurons was first achieved in a series of pioneering work
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by the Miesenböck group, by heterologously expressing an invertebrate rhodopsin with other
interacting proteins (20), and ligand-gated ion channels that can be activated by synthetic photo-
caged precursors (21,22). Around this time, microbial rhodopsins that function as single
component light-gated ion channels were discovered (23-25). Remarkably, these microbial
photoreceptors could be heterologously expressed in mammalian neurons to optically trigger
action potentials with millisecond timescale precision (6). These findings, along with the fact
that these rhodopsins contain a covalently bound all-trans retinal chromophore naturally
produced in nearly all cell types including mammalian cells and tissues (26,27), catalyzed the
wide adoption of these molecules in neuroscience.
Genetic targeting: cell-type specificity and beyond
As discussed above, cell-type specificity is a defining feature of optogenetic experiments. In
some ways, gain or loss of function experiments on distinct cell types is analogous to genetic
knock-in or knockout experiments in molecular biology. However, while the definition of gene
is clear, the definition of cell type can be ambiguous and variable. This is particularly true in
highly complex systems such as the mammalian neocortex (17,28). To account for such high
complexity, many parameters have been used to describe each cell type, such as developmental
lineage, anatomical location, dendritic and axonal morphology, electrophysiology, and gene
expression. Nonetheless, recent advances in single cell analysis methods have provided evidence
that gene expression patterns can capture many of these diverse cellular phenotypes (12,29).
Therefore, the use of genetically encoded tools is well justified for probing the function of each
cell type, and is likely to expand as we gather more gene expression data.
In principle, optogenetic tools can be driven by cell-type specific promoters or enhancers (30,31)
to achieve cell-type specific expression. However, this approach is not suitable in many cases for
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several reasons. First, it is rare to find a single genetic regulatory element exclusive for a given
cell type. Cell identity in most cases seems to be defined by a combination of multiple gene
expression patterns (10,12,17,32). Even when cell-type specific genetic regulatory elements are
accessible, they may not drive high enough expression of optogenetic tools for efficient control
(33). Moreover, reproducing endogenous expression pattern of genes requires the entire
transcription unit and all associated regulatory factors (13), which are unknown in many cases,
and can be hard to transport. A widely used strategy to achieve cell-type specific expression that
circumvents these problems is using viral vectors in transgenic animals expressing recombinase
in genetically defined population of cells. In this approach, viral vectors such as lentivirus or
adeno-associated virus (AAV) carry an optogenetic tool under relatively strong promoters such
as EF-1a promoter (Figure 1). The optogenetic tool encoding gene is initially present in reverse
frame to prevent expression, and requires recombinase activity for inversion and subsequent
expression (Figure 1). The recombinase gene expression is driven in genetically defined cell
types, for which the expression level is relatively unimportant. Cre-recombinase has been widely
used for this purpose, due to the availability of increasing number of cell type specific Cre-
transgenic lines (34). Recently, this approach has been further developed to enable targeting of
neural cell types defined by multiple genetic markers. Transgenic animals that express multiple
recombinases driven by independent promoters are infected with viral vectors carrying a
combination of these recombinase sites, resulting in intersectional targeting (35,36).
Using viral vectors also enable other useful modes of targeting. One approach is to target
neurons based on their axonal projections or synaptic connectivity (Figure 1). For example,
modified rabies virus can retrogradely transduce neurons (37,38), while herpes simplex virus and
vesicular stomatitis virus can anterogradely transduce across axon terminals (39,40). These
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viruses can be used to deliver Cre-recombinase transsynaptically (39,41), allowing the targeting
of neurons projecting to genetically defined populations. Recently, it was also shown that certain
serotypes of AAV also can mediate efficient retrograde transduction (42). These tools enable
targeting optogenetic tools based on their axonal projection (43-45) and potentially synaptic
connectivity.
Another targeting strategy is to express genes using promoters and enhancers induced by
neuronal activity. These genetic elements were identified from immediate early genes that are
known to rapidly induce expression upon neuronal activity (46,47). In many cases, these
elements are triggered by calcium influx that activates calcium-dependent kinases (48). These
promoters have been used to specifically express optogenetic tools in neurons that gain activity
during specific behavioral tasks in rodents, such as fear conditioning (49). In a set of impressive
demonstrations, Liu and others showed that these neurons could be specifically re-activated
using the light-gated ion channel channelrhodopsin-2 (ChR2) driven by the activity-dependent
promoter of c-Fos (49,50). This re-activation recreated the original behavior with only light-
activation, indicating optically controlled memory recall (49). Ramirez and others later used the
same approach to trigger fear response by optically activating a set of neurons activated during
fear conditioning, demonstrating the creation of false memory (51). Conceptually, these
demonstrations show that optogenetics can be used to reactivate cell populations specifically
activated during a behavioral task, enabling activity-based targeting. Considering the diverse
functionality of transcription factors that can sense a wide range of input signals (52-54), such
responsive transcription may be a generalizable approach to express optogenetic tools to control
cell populations that drive a specific functional output.
Customizing photoreceptors for designed control: modes of action
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The use of microbial rhodopsins in mammalian neural cells is an inspiring demonstration of
customizing naturally existing photoreceptors for controlling physiological properties in a
completely new context. Owing to the diverse modes of action found in photoreceptors (Figure
2), the concept of re-purposing naturally existing parts to control new biological functions has
been implemented in many applications, and is expected to further develop as new
photoreceptors continue to be discovered.
Microbial rhodopsins (type 1 rhodopsins) mediate either transmembrane ion transport (Figure 2a,
b) or light sensing through signal transduction (Figure 2c). Microbial rhodopsins that mediate
light-driven ion transport have been widely used to control membrane potential in mammalian
neurons (7). They can be categorized into two mechanistically distinct forms: ion pumps (Figure
2a) that can transport ions against their gradient and channels (Figure 2b) that passively conduct
ions along the gradient established by other active transporters. Interestingly, all known light-
driven ion pumps result in hyperpolarization of membrane potential through outward transport of
proton or inward transport of chloride ion (55). When heterologously expressed in mammalian
neurons, they generate enough current to enable optical silencing (7,56-58). Recently, a light-
driven chloride pump with high sensitivity for far-red light named Jaws has been described that
enabled non-invasive neural silencing through the intact mouse skull (59). A light-driven
outward sodium pump (KR2) has also been found recently (60), that may be used for optical
silencing of neurons. Unlike ion pumps, microbial rhodopsin ion channels cannot transport ions
against their gradient, but enable control of membrane potential through selective conduction of
specific ions. First described examples are channelrhodopsins found in the green alga
Chlamydomonas reinhardtii (23,24), which conduct cations including proton, sodium, potassium,
and calcium ions (24). Even though the conductance of these channels are relatively low (about
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100-fold lower than high-conductance ion channels) (61,62), their current depolarize mammalian
neurons above their threshold to initiate action potentials (6). Recently, more than 60
channelrhodopsin homologues were identified by conducting a systematic search of
transcriptome from 127 species of alga (63). This study resulted in a high sensitivity
channelrhodopsin with extremely fast channel kinetics named Chronos, and a red-sensitive
channelrhodopsin named Chrimson that as a pair enabled independent multi-color activation of
two distinct neural populations (63). Another recent study identified anion-conducting light-
gated channels in the cryptophyte Guillardia theta that enabled rapid and reversible optical
silencing of rat neurons (64). So far, microbial sensory rhodopsins have not been applied in
optogenetic experiments, perhaps due to the requirement of transmembrane or soluble
transducers that are not readily compatible with other cell types. In depth discussion of
biophysical properties of microbial rhodopsins and their impact as optogenetic tools have been
discussed elsewhere (65).
Animal rhodopsins (type II rhodopsins) share structural homology to microbial rhodopsins, but
act as G-protein coupled receptors (GPCRs) that upon light illumination, catalyze GDP to GTP
exchange in heterotrimeric G proteins. In fact, rhodopsins constitute the largest GPCR family –
over 700 rhodopsin-like GPCRs have been found in humans (66). Based on the extensive
structure-function relationship studies of GPCRs (67,68), Khorana and colleagues demonstrated
that the cytoplasmic domains of a bovine rhodopsin can be replaced by analogous sequences
from a non-light sensitive GPCR, such as β2-adrenergic receptor (β2-AR) to create a chimeric
light-sensitive β2-AR (69). This strategy has been extended to create light-driven rhodopsins
coupled to Gq and Gs signaling pathways (70), Gi/o coupled pathways (71,72). It was also found
that heterologous expression of unmodified forms of rhodopsins into non-expressing cells can
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mediate light-driven activation of endogenous G-proteins of the host cell (73). Unlike microbial
rhodopsins that keep the retinal chromophore throughout the photocycle, certain animal
rhodopsins such as bovine rhodopsin lose the chromophore after light-activation, requiring
consistent supply of retinal cofactors (74). Fortunately, in mammalian cells and in brain tissue,
sufficient concentration of retinal cofactors are present to generate enough functional rhodopsins.
However, even in cell types where the retinal chromophore is less abundant, the retinal cofactor
supplementation may be avoided by using ‘non-bleaching’ G-protein coupled opsins that remain
associated with the chromophore (75).
Other than rhodopsins based photoreceptors, light-sensing proteins found in plants and microbes
have been applied in controlling cell signaling in diverse cell types, including mammalian cells,
yeast, and bacteria. These proteins function by inducing light-triggered conformational change
coupled to other protein domains (Figure 2e), homo/hetero dimerization (Figure 2f), or
multimerization (Figure 2g). A prominent example that undergoes conformational change upon
light-activation is the Light-Oxygen-Voltage (LOV) domains found in photoreceptor systems of
plants, fungi, and bacteria (76,77). In these systems, LOV domains may control the activity of an
effector domain directly fused to it through allosteric coupling or steric inhibition. For example,
the bacterial chemosensor FixL was made light-activatable by replacing its Per-ARNT-Sims (78)
motif (which is structurally homologous to LOV domains) with the LOV domain Ytva (79). In
this example, an a-helical coiled coil linker of the LOV domain undergoes a rotational
movement upon light activation, which activates a histidine kinase domain fused to it. In other
examples, the conformational change in the LOV domain leads to unfolding of an a-helix in the
lit-state, resulting in ‘uncaging’ of the effector domain fused to it (Figure 2e). Such light-
activated uncaging approach has been successfully applied in several synthetic constructs
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including a photo-activatable GTPase Rac1 and Cdc42 (80), peptide binding motifs (81), and
tethered toxins (82). Even though such photo-uncaging approach seems to be a generally
applicable design, in many cases it requires several levels of optimization customized to each
molecule for effective control. For instance, in the photo-activatable Rac1, Ca2+-mediated
interaction between residues at the interface of the LOV domain and Rac1 turned out to be
crucial for effective light control (83). Since such conformation-dependent interactions are hard
to be predicted, de novo design of a photo-activatable construct still remains a challenge. Such
difficulties may be overcome by using Dronpa, which is a photo-activated fluorescent protein
that seems to allow modular design of optical control. Upon light-activation, Dronpa not only
changes fluorescence, but also monomerizes through the unfolding of a β-sheet (84). This feature
has been used to design photo-activatable GTPases and proteases (85).
Another mode of action found in photoreceptor domains is light-induced interaction. Certain
LOV domains, such as the fungal LOV domain Vivid (VVD) (86) and bacterial LOV domain
EL222 (87) homo-dimerize upon light activation (Figure 2f). Other photoreceptor domains such
as Arabidopsis thaliana Cryptochrome-2 (CRY2) and Phytochrome B (PhyB) hetero-dimerize
with a specific partner (Figure 2f), cryptochrome-interacting basic helix-loop-helix 1 (CIB1) and
phytochrome interacting factor (PIF), respectively (76). Such light-induced interactions have
been used primarily to control intracellular signaling, by recruiting signaling proteins to or away
from a specific intracellular site of action, resulting in signal activation or inhibition through
sequestration. Activities of various kinases, including Ras/ERK (88), phosphoinositide-3 kinase
(PI3K) (89), and receptor tyrosine kinase (90) (91) have been controlled using this approach.
Several studies also demonstrated optical control of DNA transcription using light-induced
binders that mediate the recruitment of transcription activation domains (92,93) or activation of
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Cre-recombinase (94,95). Interestingly, CRY2 has been shown to undergo multimerization upon
illumination (Figure 2g), which has been used to mediate light-dependent activation of Rho
GTPase (96), RTK fibroblast growth factor receptor (97), and actin polymerization (98). The use
of photoreceptors for optogenetic control of cellular signaling pathways has been extensively
reviewed recently (65,99,100).
Engineering photoreceptors: a multi-dimensional problem
Even though it seems that natural photoreceptor systems rely on modularity of light-activated
protein domains for controlling diverse signaling pathways (76,101), development of a new
optogenetic tool in general requires significant engineering efforts for effective control. One
major challenge is achieving adequate expression of optogenetic tools. As seen in the case of
microbial opsins, the expression level of an optogenetic construct may be one of the limiting
factor in achieving effective control. Heterologous expression of photoreceptor domains depends
on the host cell type, and may yield low levels of functionally active form of the optogenetic tool.
For example, a study that systematically compared channelrhodopsin homologues found that less
than half of them show detectable ion conduction when expressed in a mammalian cell line, and
even less are functionally active in neurons (63). One strategy to improve functional expression
is to enhance the protein trafficking to the desired compartment using targeting sequences
(58,102). However, this approach is not generalizable (103), and achieving adequate expression
of optogenetic tools in the desired cell type remains a challenge.
Another major challenge is the fact that optogenetic tools require multiple properties to be
optimized. For example, critical properties of channelrhodopsins that impact their effectiveness
in controlling neural activity include ion conductance, light sensitivity, spectral sensitivity,
channel kinetics, and ion selectivity. Structure-guided mutagenesis studies have resulted in
108
improved channelrhodopsins with fast kinetics (104), enhanced photocurrent (105,106), red-
shifted spectral sensitivity (107), and altered ion selectivity (108,109). These studies
demonstrated that key properties can be tuned using mutagenesis, but also revealed that a single
mutation often affects multiple properties (Figure 3). For example, mutations at E123 in ChR2
(corresponding to D85, which is the Schiff base counter-ion in bacteriorhodopsin), affect channel
kinetics and red-shifts the action spectra (104). Mutation H134R in ChR2 enhances photocurrent
perhaps due to improved membrane expression, but slows channel kinetics (62,110). Mutations
C128 and D156 slow down the channel kinetics drastically, which help to keep open channels for
longer periods of time, effectively enhancing the light sensitivity (111,112). The trend of
multiple property modulation by a single mutation is also found in LOV domains. For example,
in the Avena sativa phototropin 1 LOV2, mutations in the highly conserved residue Q513 reduce
the structural changes between the dark and the lit state and slow down the dark state return rate
(113). In addition, in the LOV domain VVD, mutations in M135 and M165 slow recovery
kinetics and enhance the affinity of the lit-state VVD dimer (114). Therefore, optimization of an
optogenetic tool requires multi-dimensional characterizations to measure all essential parameters
(112), and in certain cases individual properties may not be independently optimized. In other
words, the fitness landscape of photoreceptors is a multi-dimensional space composed of
potentially dependent parameters. Since typical screens used for protein engineering rely on
rapid measurement of one or two parameters, strategies such as directed evolution to optimize
one parameter may result in de-optimization of others that are critical for tool performance.
Currently, the general strategy is to combine random mutagenesis based methods with structure-
guided approaches and rely on additive beneficial effects of multiple mutations to yield an
109
optimal construct (108,109,115,116). In the future, screens that can characterize multiple
parameters may be developed to perform multi-dimensional optimizations.
Analogies to optical imaging: multi-color imaging vs. optogenetics
Another major direction in optogenetics has been to use multiple colors of light to control two
independent cell types or processes using photoreceptor domains that have distinctive spectral
sensitivities. This approach has been extensively explored using microbial opsins. For instance,
ChR2, which has maximal sensitivity for blue light has been combined with a halorhodopsin
which can be activated with yellow light to enable bi-directional control of neural activity
(56,57). Proton pumps with separation in their spectral sensitivity have been used to achieve two-
color neural silencing (58). Even though these studies demonstrated that triggering or inhibiting
action potentials in neurons can be controlled by leveraging the difference in spectral sensitivity
of microbial rhodopsins, a recent study showed that achieving multi-color control relying on
spectral separation alone may result in cross-talk, due to the inherent blue-light sensitivity found
in rhodopsins (63,82). Even the red-sensitive channelrhodopsin Chrimson was shown to trigger
action potentials under strong blue light in neurons with high expression levels (63). However,
detailed biophysical characterization of Chrimson revealed that its channel opening rate under
blue light is substantially slower than that under red light, and is also dependent on light intensity
(63). Therefore, cross-activation of neurons under blue light was minimized by pairing Chrimson
with the fast and light-sensitive channelrhodopsin Chronos and limiting the intensity and
duration of blue light (63). The blue-light sensitivity of microbial rhodopsins may not be
completely abolished unless the choromophore itself is altered. Many photoreceptors, including
LOV domains and cryptochromes are also maximally sensitive to blue light. Therefore, strategies
other than relying on spectral separation alone, such as leveraging differences in light sensitivity
110
and kinetics under blue light may be necessary to reduce cross-talk for multi-color control using
other photoreceptors.
Although the concept of multi-color control in optogenetics may seem analogous to multi-color
fluorescence imaging, there are several practical differences that require caution in designing
optogenetic experiments with multiple light sources. In multi-color fluorescence imaging, cross-
activation of fluorophores occurs quite often, but the emission from multiple fluorophores can be
filtered to obtain cross-talk free images. In experiments using light-activated proteins, any cross-
activation causes change that cannot be filtered or eliminated. Depending on the biological
question asked, small changes induced by cross-activation may become important, such as in
experiments that focus on sub-threshold changes in membrane potential. Therefore, when
designing experiments that require multiple sources of light, such as using light-gated ion
channels with calcium reporters (117), the wavelength of light used for imaging should be
chosen cautiously to prevent activation of optical actuators. It is notable that blue-sensitive
channelrhodopsins that seem to have negligible red-sensitivity can cause membrane
depolarization under intense red light (116). As described above, strategies that capitalize on all
aspects of biophysical properties of photoreceptors may be used to prevent cross-activation, but
may be difficult to completely eliminate small changes induced.
Conclusion and future perspectives
Over the past decade, optogenetics has made a major impact on neuroscience research, by
enabling the control of specific cell types in the intact brain. A powerful integration of optical
control methods with various detection methods enables closed-loop control of biological
systems, as achieved by several studies that demonstrated closed-loop control of neural circuit
(118-120) and feedback-regulated control of cell signaling (121) and gene expression levels
111
(122). These studies indicate that optogenetic approaches will enable us to reveal the underlying
mechanisms behind complex intracellular signaling systems or multi-cellular dynamics and
precisely tune it to achieve a desired outcome. They will also allow us to develop and test models
of intact, complex biological systems.
Successful development of new optogenetic tools and their implementation is an
interdisciplinary effort that requires tools tailored to the specific biological process studied.
Targeting and achieving optimal expression in the desired cell type need to be tested, and the
biophysical properties of photoreceptors used need to be optimized to the spatial and temporal
properties to be controlled. Therefore, users of optogenetic tools need to understand their
biophysical characteristics, and tool developers need to identify key properties and find ways to
tune them.
112
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