sponge… N,N,N′,N′-tetramethyl-1,8-naphthalenediamine, DIPEA… N,N-diisopropylethylamine, β-NMN… β-nicotinamide monophosphate, CDI… N,N’-carbonyldiimidazole.
Thus, 2-modified adenosine 71 was prepared from 15 and trifluoro-N-(prop-2-yn-1-yl)acetamide[157] in a Sonogashira reaction. After phosphorylation to 72 and deprotection, free amine 73 was isolated in low yields. It seems that the formation of the unreactive side
product as with 60 is even enhanced, when applying a shorter linker. Consecutive coupling of amine 73 with active ester 70 in aqueous buffer failed completely, also at pH 8.9. This is most probably due to the fact, that 70 precipitates as soon as there is water in the reaction mixture. Other amide-bond formations were tried under water-free conditions, such as in situ activations of the carboxylic acid with HBTU or pentafluorophenyl trifluoroacetate in DMF or preparing the NHS ester. Most successful was the coupling between NHS ester 70 and lyophilised 73 in DMF using DIPEA as base yielding 16% of 74. At this point, this synthesis route was stopped, because yields were too low to proceed.
Finally, the reactions were tried without shortening the linker using 6-modified phosphate 68. Waterfree-coupling now yielded 33% of compound 75. Having enough material for an NAD+ coupling in hand, the reaction was tried. Although compound 76 was successfully formed as proved by high resolution mass spectrometry, purification remained unsuccessful.
Usual ion exchange chromatography failed, because the compound strongly adsorbed to the column material unwilling to elute with different buffer and solvent mixtures. Direct HPLC purification was not able to separate unreacted 75 from 76.
In summary, all attempts to obtain BODIPY®-modified NAD+ in pure and acceptable yields failed. It seems that the combination of this hydrophobic dye and hydrophilic NAD+ is unfavourable. For future attempts to synthesise a dye-modified NAD+ with fluorophores excitable at 488 nm, I recommend using rhodamine scaffolds, e.g. Carboxy-Rhodamine 77.
Fluorescein instead possesses disadvantageous photophysical properties such as fast bleaching[137] and Atto® 488 might introduce as well synthetic problems due to two additional negative charges in the dye structure. Rhodamine 77 is structurally closest to the approved TMR scaffold and procedures are available to separate the regioisomers formed during synthesis at the level of methyl esters.
5.3 Biochemical Evaluation of NAD
+Analogues 11-12
After synthesising derivatives 11 and 12, their substrate properties were investigated using the earlier described ADP-ribosylation assays with histone H1.2 as acceptor (Figure 22A) and additionally in automodification assays[59], where ARTD1 acts as its own acceptor[37]
(Figure 22B).
When the reactions were performed in the absence of natural NAD+, the histone was only modified with fluorescent PAR chains, while using compound 11 (Figure 22A, lanes 4 vs 7).
Applying a 1:1 mixture with natural NAD+, PAR chains were also formed using 12 (lane 8).
This outcome is in line with the results obtained with the reporter-tagged NAD+ analogues in chapter 4.4. Of note, the incorporation of 2-modified derivative results in shorter PAR chains (lane 5 vs lane 2) than obtained with natural NAD+. This suggests that the modification introduced might affect polymer elongation. Interestingly, incubation in the absence of enzyme leads to some minor staining of the histone (lane 3 and 6). This might be due to electrostatic attraction between the negatively charged NAD+ and the positively charged histone and/or to non-enzymatic addition of ADP-ribose at lysine residues via Schiff base formation.[158]
To support these results, the NAD+ analogues were additionally tested in an auto(ADP-ribos)ylation assay (Figure 22B). Here, the fluorescent signal indicating auto-modified ARTD1 was also observed only, when applying 11 (lanes 3 vs 4). On the one hand, these findings suggest that the initial attachment, the polymerisation of ADP-ribose and the
Dye Modified NAD+ Analogues
elongation of already existing chains are differently affected by the introduced modifications.
On the other hand, they show that 11 competes better against the natural substrate during initiation. Whereas PAR chains formed in vitro exhibit a mean chain length of 100 ADP-ribose monomers[17], polymers synthesised in cells were found to be on average 10 units long.[19] Considering that cellular PAR is significantly shorter than in vitro, a cellular NAD+ probe must prove efficient enough in both initiation and elongation reactions. Since these criteria are better met by 11, experiments were continued using this analogue only.
Figure 22. SDS PAGE analyses of ADP-ribosylation of histone H1.2 with ARTD1 (A) and ARTD1 automodification (B) using NAD+ analogues 11-12. Upper panel shows Coomassie Blue staining;
lower panel shows TMR fluorescence. Total concentration of NADs was 1 mM. Controls were performed using either natural substrate (A: lane 1 and 2, B: lane 2) or A: no enzyme (lane 1, 3 and 6) or B: loading the same amount of ARTD1 (lane 1).
As a next step, compound 11 was studied in an ADP-ribosylation assay of H1.2 using different ratios of modified to unmodified NAD+. As seen in Figure 23, PAR formation can be detected within a wide range of ratios. At ratios of modified NAD+ to unmodified NAD+ below 1:5, almost no differences in polymer formation are visible in vitro (lanes 6 to 10 vs 11). This demonstrates that compound 11 is a competitive substrate for PARylation.
Figure 23. SDS PAGE analysis of ADP-ribosylation of histone H1.2 with ARTD1 using NAD+ analogue 11 and natural NAD+. Upper panel shows Coomassie Blue staining; lower panel shows TMR fluorescence.
The total concentration of NADs was 1 mM with different ratios of modified to unmodified NAD+ as indicated. Controls were performed using either natural substrate (lane 1 and 11) or no enzyme (lane 1 and 2).
5.4 Towards A Cellular Application
Based on in vitro data, the next step was to translate these results into the setting of a living cell. As already mentioned in chapter 4.6, I collaborated with Annette Buntz from the group of Prof. Dr. Zumbusch for the cellular applications. She performed the NAD+ transfer methods described in here.
The major challenge for cellular applications was to transfer the negatively charged NAD+ molecule through the cell membrane into the cytosol, where it is available for enzymatic consumption. Moreover, cells need to remain viable and to the greatest possible extent undamaged for further biological manipulations.
Different methods are known to help charged molecules passing the cell membrane.[159] In the case of nucleotides, electroporation,[160] transient permeabilisation[116] and the carrier peptide Pep-1[55] have been applied previously. During electroporation, holes are introduced into the membrane by manipulating the electronic potential with the help of electrodes[160] and thus allowing the diffusion of small NAD+ probes into the cytoplasm. Optimising the electronic parameters for the cell type and application, cells are known to be structurally intact and viable for a certain time. Within transient permeabilisation (Chapter 4.6), small detergent concentrations are responsible for the same effect. The carrier peptide Pep-1 was developed to deliver peptides, proteins and even antibodies into cells in a non-toxic, endocytosis-independent manner by forming non-covalent interaction with its cargo.[161] The amphipathic peptide Pep-1 comprises a tryptophan-rich so-called ‘hydrophobic’ domain, a hydrophilic domain derived from a nuclear localisation signal (NLS) of simian virus 40 (SV40) large T-antigen, and a spacer between them.
Firstly, all three methods were tested and their handling was optimised. In Figure 24, representative confocal images of cell samples treated either with electroporation, permeabilisation or carrier-peptide and compound 11 are depicted. Comparing qualitatively the fluorescence signal, electroporation was found to be the least effective technique.
Moreover, when comparing the transmission pictures, the use of carrier-peptide resulted in
Dye Modified NAD+ Analogues
healthy looking and structurally intact cells, whereas the permeabilisation method already showed first apoptotic bodies inside of cells. These bodies indicate that the treated cells will soon undergo apoptosis and will not be viable for prolonged microscopic studies. In case of the cellular assay performed in chapter 4.6, the cells were immediately damaged and fixed, making the permeabilisation a suitable method. However, for the anticipated real-time experiments, cells need to be viable ideally for several hours. Thus, we decided to use the carrier peptide method for further optimisations.
Figure 24. Comparing methods for the uptake of NAD+ analogue 11 into HeLa cells. Electroporation: 160 V/cm, 0.4 cm electrodes, 100 µs pulses, NAD+ concentration: 100 µM. Transient permeabilisation: Triton-X-100 0.01%, 4 °C, 12 min, NAD+ concentration: 100 µM. Carrier peptide Pep-1: 100 µM Pep-1, NAD+ concentration: 25 µM,1 h at 37 °C. Scale bar: 50 µm.
Next, different cargo-carrier ratios were tested and the labelling efficiency was analysed by flow cytometry. In addition, cell viability was estimated by staining dead cells with Sytox Blue®. Incubating cells for one hour with 25 µM of 11 and 100 µM Pep-1, more than 90% of the cells were efficiently labelled and cell viability was not affected at all applied concentrations (Figure 25). Accordingly, these conditions were used in all following cell experiments.
Figure 25. Flow cytometric analysis of uptake efficiency and cell viability after treatment with NAD+ analogue 11.
HeLa cells were treated with 11 (0.5-25 µM) and carrier peptide Pep-1 (5-100 µM) for 1 h.
Subsequently, the cells were harvested and analysed by flow cytometry. Dead cells were additionally labelled by Sytox Blue. (A) Representative image of flow cytometry data to illustrate gating strategy of cell populations. Numbers indicate percentages of the respective quadrants. (B) Fluorescence intensities were measured from viable (i.e. Sytox Blue-negative), TMR-positive cells. (C) The fraction of TMR-positive and TMR-negative living cells was determined. Three independent experiments were performed. Error bars represent standard errors of the mean (SEM).
5.5 Live-Cell Monitoring of Poly(ADP-Ribos)ylation
The main aim of this thesis was to visualise the formation of PAR inside of a living cell and in real-time. After developing a dye-labelled NAD+ analogue, proving its performance in vitro and finding a suitable method to introduce it into cells, we started the collaboration with the Lab of Prof. Dr. Elisa Ferrando-May at the University of Konstanz. All experiments performed with their experimental setup were conducted by a fellow PhD student Eva Gwosch. Annette Buntz and I assisted in sample preparation, execution and data analysis.
For monitoring PAR formation at sites of DNA damage, a confocal microscope equipped with a femtosecond fibre laser source was used. The microirradiation of cell nuclei with femtosecond laser pulses of 775 nm results in DNA damage from three-photon excitation equivalent to a wavelength of 258 nm.[162] The photodamage is spatially confined in three dimension to a volume corresponding to the focal spot and leads to the activation of ARTD1[163] and other proteins associated to DNA repair. After cells were loaded with NAD+ analogue 11 with the help of carrier-peptide Pep-1, cell nuclei were irradiated along a line track (Figure 26A). Confocal fluorescence images were recorded every 10 seconds at a wavelength of 555 nm. In this way, cellular redistribution of the TMR-fluorescence signal corresponding to PAR formation was monitored (Figure 26B). Performing irradiation on 30
Dye Modified NAD+ Analogues
different cells and in three independent experiments, image data could additionally be quantified. Quantification was carried out by determining the enhancement of the fluorescence signal for each time point of the irradiated region of interest relative to the surrounding area with the help of an ImageJ macro developed for this purpose (Figure 26C).
Figure 26. Live-Cell monitoring of PARylation upon DNA damage by microirradiation using NAD+ analogue 11.
(A) Scheme illustrating the experimental setup. HeLa cells were loaded with 11 and microirradiated for DNA damage. (B) Quantitative analysis of relative enhancement of fluorescence signal at DNA damage site compared to the surrounding area (n=32 cells). Control experiments were performed with 10 µM ARTD inhibitor ABT888 (n=28 cells). Three independent experiments were performed.
Error bars represent standard errors of the mean (SEM). (C) Exemplary confocal microscopy image sequence. Scale bars: 7 µm.
Within seconds after the laser pulse, a strong increase in fluorescence intensity was observed at the site of DNA damage (see movie on attached disk). For control, the same experiment was performed in the presence of the ARTD inhibitor ABT888.[164] In this setup, olaparib cannot be used, because pre-incubation with this inhibitor results already in DNA damage caused by ARTD trapping.[165] Applying the inhibitor, fluorescence intensity enhancement along the track was completely abrogated. This indicates that the accumulation of 11 and its processing at the site of damage can be attributed to ARTD1 activity.
Surprisingly, the strong fluorescence increase corresponding to PAR formation is additionally followed by a slower fluorescence intensity decay starting approximately after 50 seconds. This indicates that the fluorescent PAR chains are furthermore degraded by PAR glycohydrolases. Thus, not only PAR formation was recorded in real-time, but a whole PAR turnover was monitored. The approach developed in here enables to study cellular, stimuli-dependent PARylation dynamics in real-time and should thus be a benefit to the PAR research community.
Despite these positive results, the application of carrier-peptide Pep-1 displayed also a limitation. Approximately one hour after removing the transfection mix, cells under the microscope went into apoptosis. Control experiments applying only the carrier peptide indicated, that this apoptosis is due to the transfection method and not due to the NAD+ analogue. This downside was overcome by preparing multiple sample cells with one hour intervals. Using less peptide (25 µM) delayed this process for half an hour, but cells are not
as intensively labelled. As conclusion, a better and milder delivery technique would be highly desirable.
Initially, compound 12 was also tested in this setup and no fluorescence enhancement has been observed after DNA damage by laser (Figure 27). Although 12 was found to produce PAR in mixtures with natural NAD+ in vitro, its modification seems to be sterically too demanding to be useful for in vivo approaches. This might be an explanation, why the commercially available 6-modified NAD+ analogues such as 6-biotin-NAD+ have not been widely used for studying ADP-ribosylation processes in more complex systems such as cells.
Figure 27. NAD+ analogue 12 is not recruited to DNA damage sites. 100 µM of 12 were delivered into HeLa cells and irradiated as described above. Cells were imaged with a confocal fluorescence microscope. An exemplary confocal image sequence is shown and no fluorescence enhancement at DNA damage site was observed. Scale bars: 7 µm.
5.6 Fluorescence Lifetime Imaging of Protein-Specific PARylation in Cells
As most of the functions of PARylation are mediated through covalent and non-covalent binding of proteins, it would be as well of great benefit to study these interactions on a protein-specific level. For this purpose, proximity-based FRET approaches have been very useful. One of the most robust methods to determine FRET is the measurement of fluorescence lifetimes.[100] Within her PhD thesis, Annette Buntz constructed a so-called fluorescence lifetime imaging microscope (FLIM) being able to resolve spatially fluorescence lifetime changes in the dimension of living cells.[166] Thus, we opted to implement the new NAD+ analogue 11 in combination with eGFP-tagged proteins into this FLIM-FRET technique and to detect ADP-ribosylation in a protein-specific manner as it has already been useful for other posttranslational modifications.[167-168] All in vitro experiments were performed and optimised by Annette Buntz and are described in detail in her PhD Thesis.[166] For the final cell-based experiments described in the following, Eva Gwosch and I assisted in sample preparation and execution. The recombinant purified eGFP-tagged ARTD1 was provided by Sascha Beneke, University of Konstanz. The plasmid coding for eGFP-tagged ARTD1 and macroH2A was already available in the group of Prof. Dr. Ferrando-May.
First and for proof-of principle, the automodification of ARTD1 was chosen as an example for the detection of a covalent PAR interaction. For this purpose, ARTD1 was fused to the FRET donor eGFP, while the TMR label present on the NAD+ analogue functions as FRET acceptor. This FRET pair has a calculated Förster Radius of 5.8 nm and can effectively be used to detect donor-acceptor distances of 2 to 10 nm. To confirm that the eGFP-tag does not impair ARTD1 activity, wild type ARTD1 and eGFP-ARTD1 were compared in the usual ADP-ribosylation assays (Figure 28) indicating no different in vitro behaviour.
Dye Modified NAD+ Analogues
Figure 28. SDS PAGE analysis comparing wild type ARTD1 (wt) with eGFP variant in ADP-ribosylation of histone H1.2 (A) and automodification of ARTD1 (B). Left panel shows Coomassie Blue staining; right panel shows TMR-fluorescence signals. The total concentration of NADs was 1 mM. (A) Reactions were performed with natural NAD+ (lane 1 and 4), with a 9:1 mixture of natural NAD+ and NAD+ analogue 11 (lane 2 and 5), without enzyme (lane 3) or with ARTD inhibitor olaparib (I, lane 6). (B) As a control, the same amount of ARTD1 was loaded (lane 1 and 5). Reactions were performed with natural NAD+ (lane 2 and 6), with a 9:1 mixture of natural NAD+ and NAD+ analogue 11 (lane 3 and 7), without enzyme (lane 4) or with ARTD inhibitor olaparib (I, lane 8).
When eGFP-ARTD1 is modified with TMR-containing ADP-ribose, FRET can be detected by a reduction in the fluorescence lifetime of the eGFP-label, whereas modification with natural NAD+ has no effect on the lifetime (Figure 29A). Annette performed a set of in vitro experiments, where consecutive modification of eGFP-ARTD1 with multiple TMR-labelled ADP-ribose units led to a prominent and steady decrease of the eGFP fluorescence lifetime, indicative of increasing FRET. She used either the purified protein or lysates of HEK293T cells expressing eGFP-ARTD1 and monitored PARylation of ARTD1 in real-time. The effect was not observed, when the reaction was performed in the absence of dsDNA or upon addition of ARTD-inhibitor olaparib[149].
After successful in vitro measurements, cellular imaging was advanced. For this purpose, HeLa cells were transfected with the eGFP-ARTD1 plasmid overnight and transiently expressed the protein the next day. Then, these cells were treated with the delivery mix containing 11 and Pep-1 as described earlier (Chapter 5.4). Unfortunately, this double treatment stressed the HeLa cells and it was difficult to find doubly labelled, viable cells.
However, sufficient numbers of cells could be subjected to laser microirradiation as described earlier (Chapter 5.5) by being fast and preparing multiple cell samples. Laser-induced DNA damage led to the enrichment of both eGFP and TMR fluorescence signals at the irradiated sites, indicating that ARTD1 is recruited and fluorescent PAR is formed (Figure 30A). After two to three minutes after laser damage, the cells were fixed and subsequently analysed on the FLIM microscope (Figure 29B). The fluorescence lifetimes of eGFP-ARTD1 were measured for every image pixel and is displayed via intensity-weighted
pseudo-colours (e.g. pixels with higher intensity appear brighter). At DNA damage sites, the eGFP fluorescence signals are very bright due to ARTD1 recruitment. When no fluorescently labelled NAD+ was applied, the eGFP fluorescence lifetime was found to be invariant throughout the nucleus. With 11 being present, regions of high eGFP intensity show a significantly reduced fluorescence lifetime of the donor caused by FRET between eGFP-ARTD1 and the TMR-labelled PAR. This observation is confirmed by a quantitative analysis of fluorescence lifetimes measured in 15 cells (Figure 29C).
Figure 29. Fluorescence lifetime-based read-out of protein-specific PARylation. Left: Detection of covalent ARTD1 automodification. (A) Activation of ARTD1 leads to FRET between eGFP and the TMR-label on PAR, measured by a decrease of the eGFP fluorescence lifetime. (B) HeLa cells expressing eGFP-ARTD1 were treated with 11 or left untreated, microirradiated for DNA damage induction, and fixed. Fluorescence lifetime images of eGFP-ARTD1 are intensity-weighed and presented in pseudo-colours. Scale bars: 7 µm. (C) Quantitative analysis of fluorescence lifetimes from 15 cells transfected with eGFP-ARTD1. Right: Detection of the non-covalent interaction of macroH2A-eGFP with PAR. (D) Activation of ARTD1 leads to PARylation and recruitment of macroH2A. Non-covalent interaction results in FRET between eGFP and the TMR-label on PAR, measured by a decrease of the eGFP fluorescence lifetime. (E) Fluorescence lifetime images of macroH2A-eGFP upon DNA photodamage. (E) Quantitative analysis of fluorescence lifetimes from 15 cells transfected with macroH2A-eGFP. (C), (E) Mean values ± SEM are depicted. Statistical significance was assessed with a Two-Way-ANOVA and Bonferroni posttest. The level of significance is given with n.s. … not significant, *** p < 0.001.
Dye Modified NAD+ Analogues
Figure 30. Enrichment of eGFP-fusion proteins and TMR fluorescence signals at sites of microirradiation. HeLa
Figure 30. Enrichment of eGFP-fusion proteins and TMR fluorescence signals at sites of microirradiation. HeLa